Laboratory Manual

Laboratory Research in Environmental Engineering Laboratory Manual

Monroe L. Weber-Shirk Leonard W. Lion James J. Bisogni, Jr. Cornell University School of Civil and Environmental Engineering Ithaca, NY 14853

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CEE 453: Laboratory Research in Environmental Engineering Spring 2001

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Laboratory Research in Environmental Engineering Laboratory Manual

Monroe L. Weber-Shirk Instructor [email protected] Leonard W. Lion Professor [email protected] James J. Bisogni, Jr. Associate Professor [email protected] School of Civil and Environmental Engineering Cornell University Ithaca, NY 14853 Fifth Edition

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© Cornell University 2001

Educational institutions may use this text freely if the title/author page is included.

We request that instructors who use this text notify one of the authors so that the

dissemination of the manual can be documented and to ensure receipt of future

editions of this manual.

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Table of Contents

Table of Contents

Table of Contents ...............................................................................................................5

Preface.................................................................................................................................9

Laboratory Safety ............................................................................................................10 Introduction....................................................................................................................10 Personal Protection ........................................................................................................10 Laboratory Protocol .......................................................................................................12 Use of Chemicals ...........................................................................................................13 References ......................................................................................................................19 Questions ........................................................................................................................19

Laboratory Measurements and Procedures..................................................................20 Introduction....................................................................................................................20 Theory............................................................................................................................20 Experimental Objectives................................................................................................22 Experimental Methods ...................................................................................................22 Prelab Questions ............................................................................................................24 Questions ........................................................................................................................25 Data Sheet ......................................................................................................................27 Lab Prep Notes...............................................................................................................29

Reactor Characteristics ...................................................................................................30 Introduction....................................................................................................................30 Reactor Classifications ...................................................................................................30 Reactor Modeling...........................................................................................................30 Mass Conservation.........................................................................................................34 Conductivity Measurements ..........................................................................................35 Procedures ......................................................................................................................36 Prelab Questions ............................................................................................................39 Data Analysis .................................................................................................................39 Lab Prep Notes...............................................................................................................41

Acid Precipitation and Remediation of Acid Lakes......................................................43 Introduction....................................................................................................................43 Experimental Objectives................................................................................................47 Experimental Apparatus .................................................................................................48 Experimental Procedures ...............................................................................................48 Prelab Questions ............................................................................................................53 Data Analysis .................................................................................................................53 Questions ........................................................................................................................54 References ......................................................................................................................54 Lab Prep Notes...............................................................................................................55

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Measurement of Acid Neutralizing Capacity ................................................................57 Introduction....................................................................................................................57 Theory............................................................................................................................57 Procedure .......................................................................................................................60 Prelab Questions ............................................................................................................61 Questions ........................................................................................................................61 References ......................................................................................................................62 Lab Prep Notes...............................................................................................................63

Phosphorus Determination using the Colorimetric Ascorbic Acid Technique ..........64 Introduction....................................................................................................................64 Experimental Objectives................................................................................................66 Experimental Procedures ...............................................................................................66 Prelab Questions ............................................................................................................67 Data Analysis .................................................................................................................67 Questions ........................................................................................................................68 References ......................................................................................................................68 Lab Prep Notes...............................................................................................................69

Soil Washing to Remove Mixed Wastes.........................................................................70 Objective ........................................................................................................................70 Introduction....................................................................................................................70 Theory............................................................................................................................71 Apparatus .......................................................................................................................77 Experimental Procedures ...............................................................................................78 Prelab Questions ............................................................................................................82 Data Analysis .................................................................................................................82 References ......................................................................................................................82 Lab Prep Notes...............................................................................................................85

Oxygen Demand Concepts and Dissolved Oxygen Sag in Streams .............................87 Introduction....................................................................................................................87 Theory............................................................................................................................87 Streeter Phelps Equation Development .........................................................................88 Zero Order Kinetics .......................................................................................................92 Experimental Objectives................................................................................................93 Experimental Methods ...................................................................................................94 Prelab Questions ............................................................................................................95 Data Analysis .................................................................................................................96 References ......................................................................................................................97 Lab Prep Notes...............................................................................................................98

Methane Production from Municipal Solid Waste .....................................................100 Introduction..................................................................................................................100 Theory..........................................................................................................................100 Experiment description................................................................................................110 Experimental methods..................................................................................................112

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Table of Contents

Prelab questions ...........................................................................................................114 Data analysis ................................................................................................................114 References ....................................................................................................................115 Lab Prep Notes.............................................................................................................117

Volatile Organic Carbon Contaminated Site Assessment ..........................................119 Introduction..................................................................................................................119 Experiment Description ...............................................................................................119 Experimental Procedures .............................................................................................120 Procedure (short version) .............................................................................................122 Prelab Questions ..........................................................................................................123 Data Analysis ...............................................................................................................123 References ....................................................................................................................123 Lab Prep Notes.............................................................................................................124

Volatile Organic Carbon Sorption to Soil ...................................................................126 Introduction..................................................................................................................126 Theory..........................................................................................................................126 Analysis of the Unsaturated Distribution Coefficient ( S

GK ) ........................................131

Analysis of the Saturated Distribution Coefficient ( SLK ) ............................................133

Experimental procedures..............................................................................................135 Procedure (short version) .............................................................................................136 Prelab Questions ..........................................................................................................137 Data Analysis ...............................................................................................................137 References ....................................................................................................................138 Additional References Relevant to Data Reduction ....................................................139 Symbol List..................................................................................................................140 Lab Prep Notes.............................................................................................................141

Enhanced Filtration.......................................................................................................142 Introduction..................................................................................................................142 Theory..........................................................................................................................142 Previous Research Results ...........................................................................................144 Filter Performance Evaluation.....................................................................................145 Experimental Objectives..............................................................................................145 Experimental Methods .................................................................................................145 Prelab Questions ..........................................................................................................147 Data Analysis ...............................................................................................................147 Questions for Discussion .............................................................................................148 References ....................................................................................................................148 Lab Prep Notes.............................................................................................................149

Gas Transfer...................................................................................................................150 Introduction..................................................................................................................150 Theory..........................................................................................................................150 Experimental Objectives..............................................................................................152

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Experimental Methods .................................................................................................153 Prelab Questions ..........................................................................................................154 Data Analysis ...............................................................................................................154 References ....................................................................................................................155 Lab Prep Notes.............................................................................................................156

Instrument Instructions .................................................................................................157 Compumet software .....................................................................................................157 pH Probe Calibration ...................................................................................................157 pH Probe Storage .........................................................................................................158 Procedure for Cleaning pH Gel-Filled Polymer Electrode ..........................................158 Dissolved Oxygen Probe..............................................................................................158 Gas Chromatograph .....................................................................................................160 UV-Vis Spectrophotometer..........................................................................................160

Index................................................................................................................................161

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Preface

Preface Continued leadership in environmental protection requires efficient transfer of

innovative environmental technologies to the next generation of engineers. Responding to this challenge, the Cornell Environmental Engineering faculty redesigned the undergraduate environmental engineering curriculum and created a new senior-level laboratory course. This laboratory manual is one of the products of the course development. Our goal is to disseminate this information to help expose undergraduates at Cornell and at other institutions to current environmental engineering problems and innovative solutions.

A major goal of the undergraduate laboratory course is to develop an atmosphere where student understanding will emerge for the physical, chemical, and biological processes that control material fate and transport in environmental and engineered systems. Student interest is piqued by laboratory exercises that present modern environmental problems to investigate and solve.

The experiments were designed to encourage the process of “learning around the edges.” The manifest purpose of an experiment may be a current environmental problem, but it is expected that students will become familiar with analytical methods in the course of the laboratory experiment (without transforming the laboratory into an exercise in analytical techniques). It is our goal that students employ the theoretical principles that underpin the environmental field in analysis of their observations without transforming the laboratories into exercises in process theory. As a result, students can experience the excitement of addressing a current problem while coincidentally becoming cognizant of relevant physical, chemical, and biological principles.

At Cornell, student teams of two or three carry out the exercises, maximizing the opportunity for a hands-on experience. Each team is equipped with modern instrumentation as well as experimental reactor apparatus designed to facilitate the study of each topic.

Computerized data acquisition and instrument control are used extensively to make it easier for students to learn how to use new instruments and to eliminate the drudgery of manual data acquisition. Software was developed at Cornell to use computers as virtual instruments that interface with a pH meter/ion (Accumet 50), gas chromatograph (HP 5890A), UV-Vis Spectrophotometer (HP 8452) This code is available at the course web site.

The development of this manual and the accompanying course would not have been possible without funds from the National Science Foundation, the DeFrees Family Foundation, the Procter and Gamble Fund, the School of Civil and Environmental Engineering and the College of Engineering at Cornell University.

Monroe L. Weber-Shirk Leonard W. Lion James J. Bisogni, Jr. Ithaca, NY December 22, 2000

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Laboratory Safety

Introduction Safety is a collective responsibility that requires the full cooperation of everyone in

the laboratory. However, the ultimate responsibility for safety rests with the person actually carrying out a given procedure. In the case of an academic laboratory, that person is usually the student. Accidents often result from an indifferent attitude, failure to use common sense, or failure to follow instructions. Each student should be aware of what the other students are doing because all can be victims of one individual's mistake. Do not hesitate to point out to other students that they are engaging in unsafe practices or operations. If necessary, report it to the instructor. In the final assessment, students have the greatest responsibility to ensure their own personal safety.

This guide provides a list of do's and don'ts to minimize safety and health problems associated with experimental laboratory work. It also provides, where possible, the ideas and concepts that underlie the practical suggestions. However, the reader is expected to become involved and to contribute to the overall solutions. The following are general guidelines for all laboratory workers: 1) Follow all safety instructions carefully. 2) Become thoroughly acquainted with the location and use of safety facilities such

as safety showers, exits and eyewash fountains. 3) Become familiar with the hazards of the chemicals being used, and know the

safety precautions and emergency procedures before undertaking any work. 4) Become familiar with the chemical operations and the hazards involved before

beginning an operation.

Personal Protection

Eye Protection All people in the laboratory including visitors must wear eye protection at all times,

even when not performing a chemical operation. Wearing of contact lenses in the laboratory is normally forbidden because contact lenses can hold foreign materials against the cornea. Furthermore, they may be difficult to remove in the case of a splash. Soft contact lenses present a particular hazard because they can absorb and retain chemical vapors. If the use of contact lenses is required for therapeutic reasons fitted goggles must also be worn. In addition, approved standing shields and face shields that protect the neck and ears as well as the face should be used when appropriate for work at reduced pressure or where there is a potential for explosions, implosions or splashing. Normal prescription eyeglasses, though meeting the Food and Drug Administration's standards for shatter resistance, do not provide appropriate laboratory eye protection.

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Laboratory Safety

Clothing Clothing worn in the laboratory should offer protection from splashes and spills,

should be easily removable in case of accident, and should be at least fire resistant. Nonflammable, nonporous aprons offer the most satisfactory and the least expensive protection. Lab jackets or coats should have snap fasteners rather than buttons so that they can be readily removed.

High-heeled or open-toed shoes, sandals, or shoes made of woven material should not be worn in the laboratory. Shorts, cutoffs and miniskirts are also inappropriate. Long hair and loose clothing should be constrained. Jewelry such as rings, bracelets, and watches should not be worn in order to prevent chemical seepage under the jewelry, contact with electrical sources, catching on equipment, and damage to the jewelry.

Gloves Gloves can serve as an important part of personal protection when they are used

correctly. Check to ensure the absence of cracks or small holes in the gloves before each use. In order to prevent the unintentional spread of chemicals, gloves should be removed before leaving the work area and before handling such things as telephones, doorknobs, writing instruments, computers, and laboratory notebooks. Gloves may be reused, cleaned, or discarded, consistent with their use and contamination.

A wide variety of gloves is available to protect against chemical exposure. Because the permeability of gloves of the same or similar material varies from manufacturer to manufacturer, no specific recommendations are given here. Be aware that if a chemical diffuses through a glove, that chemical is held against the worker's hand and the individual may then be more exposed to the chemical than if the glove had not been worn.

Personal Hygiene Everyone working in a chemistry laboratory should be aware of the dangers of

ingesting chemicals. These common sense precautions will minimize the possibility of such exposure: 1) Do not prepare, store (even temporarily), or consume food or beverages in any

chemical laboratory. 2) Do not smoke in any chemical laboratory. Additionally, be aware that tobacco

products in opened packages can absorb chemical vapors. 3) Do not apply cosmetics in a laboratory. 4) Wash hands and arms thoroughly before leaving the laboratory, even if gloves

have been worn. 5) Wash separately from personal laundry, lab coats or jackets on which chemicals

have been spilled. 6) Never wear or bring lab coats or jackets into areas where food is consumed. 7) Never pipette by mouth. Always use a pipette aid or suction bulb.

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Laboratory Protocol The chemistry laboratory is a place for serious learning and working. Horseplay

cannot be tolerated. Variations in procedures including changes in quantities or reagents may be dangerous. Such alterations may only be made with the knowledge and approval of the instructor.

Housekeeping In the laboratory and elsewhere, keeping things clean and neat generally leads to a

safer environment. Avoid unnecessary hazards by keeping drawers and cabinets closed while working. Never store materials, especially chemicals, on the floor, even temporarily. Work spaces and storage areas should be kept clear of broken glassware, leftover chemicals and scraps of paper. Keep aisles free of obstructions such as chairs, boxes and waste receptacles. Avoid slipping hazards by keeping the floor clear of ice, stoppers, glass beads or rods, other small items, and spilled liquids. Use the required procedure for the proper disposal of chemical wastes and solvents.

Cleaning Glassware Clean glassware at the laboratory sink or in laboratory dishwashers. Use hot water,

if available, and soap or other detergent. If necessary, use a mild scouring powder. Wear appropriate gloves that have been checked to ensure that no holes are present. Use brushes of suitable stiffness and size. Avoid accumulating too many articles in the cleanup area. Usually work space around a sink is limited and piling up dirty or cleaned glassware leads to breakage. Remember that the turbid water in a sink may hide a jagged edge on a piece of broken glassware that was intact when put into the water. A pair of heavy gloves may be useful for removing broken glass, but care must be exercised to prevent glove contamination. To minimize breakage of glassware, sink bottoms should have rubber or plastic mats that do not block the drains.

Avoid the use of strong cleaning agents such as nitric acid, chromic acid, sulfuric acid, strong oxidizers, or any chemical with a "per" in its name (such as perchloric acid, ammonium persulfate, etc.) unless specifically instructed to use them, and then only when wearing proper protective equipment. A number of explosions involving strong oxidizing cleaning solutions, such as chromic sulfuric acid mixtures, have been reported. The use of flammable solvents should be minimized and, when they are used, appropriate precautions must be observed.

Unattended Operation of Equipment Reactions that are left to run unattended overnight or at other times are prime

sources for fires, floods and explosions. Do not let equipment such as power stirrers, hot plates, heating mantles, and water condensers run overnight without fail-safe provisions and the instructor's consent. Check unattended reactions periodically. Always leave a note plainly posted with a phone number where you and the instructor can be reached in case of emergency. Remember that in the middle of the night, emergency personnel are entirely dependent on accurate instructions and information.

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Laboratory Safety

Fume Hoods and Ventilation A large number of common substances present acute respiratory hazards and

should not be used in a confined area in large amounts. They should be dispensed and handled only where there is adequate ventilation, such as in a hood. Adequate ventilation is defined as ventilation that is sufficient to keep the concentration of a chemical below the threshold limit value or permissible exposure limit.

If you smell a chemical, it is obvious that you are inhaling it. However, odor does not necessarily indicate that a dangerous concentration has been reached. By contrast, many chemicals can be present at hazardous concentrations without any noticeable odor.

Refrigerators Chemicals stored in refrigerators should be sealed, double packaged if possible,

and labeled with the name of the material, the date placed in the refrigerator, and the name of the person who stored the material A current inventory should be maintained. Old chemicals should be disposed of after a specified storage period. Household refrigerators should not be used for chemical storage.

If used for storage of radioactive materials, a refrigerator should be plainly marked with the standard radioactivity symbol and lettering, and routine surveys should be made to ensure that the radioactive material has not contaminated the refrigerator.

Food should never be stored in a refrigerator used for chemical storage. These refrigerators should be clearly labeled "No Food". Conversely food refrigerators, which must be always outside of, and away from, the chemical work area, should be labeled "Food Only—No Chemicals".

Radioactive Materials Radioactive materials are used in the Environmental Engineering laboratories.

Doors of rooms containing radioactive materials are clearly labeled. Areas where radioactive materials are used are clearly delineated with labeling tape and signs. All equipment within areas labeled radioactive are potentially contaminated and should not be touched or removed. Do not place anything into or take anything from an area labeled radioactive.

Working Alone Avoid working alone in a building or in a laboratory.

Use of Chemicals Before using any chemical you need to know how to safely handle it. The safety

precautions taken are dependent on the exposure routes and the potential harmful effects.

Routes of Exposure

1) ingestion 2) inhalation 3) absorbed through skin

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4) eye contact Each potential exposure route requires different precautions. Chemical exposure

may have acute (immediate, short term) or chronic (long term potentially cumulative) affects. Information on health hazards can be found on chemical labels and in Material Safety Data Sheets.

Material Safety Data Sheets MSDS sheets for most chemicals used in the laboratory are located on the

bookshelf in the entrance hallway of the Environmental Laboratory. Electronic versions (potentially more current) can be found using the world wide web at: http://www.cee.cornell.edu/safety/

MSDS provide extensive information on safe handling, first aid, toxicity, etc. Following is a list of terms used in MSDS:

TLV—Threshold Limit Value—are values for airborne toxic materials that are to be used as guides in control of health hazards. They represent concentrations to which nearly all workers (workers without special sensitivities) can be exposed to for long periods of time without harmful effect. TLV's are usually expressed as parts per million (ppm). TLV's are also expressed as mg of dust or vapor/m3 of air.

TDLo—Toxic Dose Low—the lowest dose of a substance introduced by any route, other than inhalation, over any given period of time and reported to produce any toxic effect in humans or to produce carcinogenic, neoplastigenic, or teratogenic effects in animals or humans.

TCLo—Toxic Concentration Low—the lowest concentration of a substance in air to which humans or animals have been exposed for any given period of time and reported to produce any toxic effect in humans or to produce carcinogenic, neoplastigenic, or teratogenic effects in animals or humans.

TDLo—Lethal Dose Low—the lowest dose (other than LD50) of a substance introduced by any route, other than inhalation, over any given period of time in one or more divided portions and reported to have caused death in humans or animals.

LD50—Lethal Dose Fifty—a calculated dose of a substance that is expected to cause the death of 50% of an entire defined experimental animal population. It is determined from the exposure to the substance by any route other than inhalation of a significant number from that population.

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Laboratory Safety

LCLo—Lethal Concentration Low—the lowest concentration of a substance in air, other than LC50, that has been reported to have caused death in humans or animals. The reported concentrations may be entered for periods of exposure that are less than 24 hours (acute) or greater than 24 hours (subacute and chronic).

LC50—Lethal Concentration Fifty—a calculated concentration of a substance in air, exposure to which for a specified length of time is expected to cause the death of 50% of an entire defined experimental animal population. It is determined from the exposure to the substance of a significant number from that population.

Chemical Labels All chemicals must be labeled. Unlabeled containers of mystery chemicals or

chemical solutions are a nightmare for disposal as well as a potential safety hazard. The OSHA Hazard Communication Standard and the OSHA Lab Standard have specific requirements for the labeling of chemicals. In a laboratory covered under the Lab Standard, if a chemical is designated as a hazardous material, that is having the characteristics of corrosivity, ignitability, toxicity (generally meaning a highly toxic material with an LD50 of 50 mg/kg or less), reactivity, etc., and if it is made into a solution or repackaged as a solid or liquid in a concentration greater than 1% (0.1% for a carcinogen) it needs to have a so called Right-To-Know (RTK) label that duplicates the hazard warnings, precautions and first aid steps found on the original label. All other chemicals must have at minimum a label with chemical name, concentration, and date prepared. "Right to Know Labels" will be made available for your use when necessary.

Table 1. NFPA hazard code ratings.

Code Health Fire Reactivity 4

Very short exposure can cause death or

major residual injury

Will rapidly or completely

vaporize at normal pressure and temperature

Capable of detonation or explosive reaction at

normal temperatures and pressures

3

Short exposure can cause serious temporary or

residual injury

Can be ignited under almost all

ambient temperatures

Capable of detonation or explosive reaction buy requires a strong

initiating source or must be heated under confinement before

initiation 2

Intense or continued exposure can cause

temporary incapacitation or possible residual

injury

Must be moderately heated or exposed to high temperature before

ignition

Undergoes violent chemical change at

elevated temperatures and pressures or reacts

violently with water.

1

Can cause irritation but only minor residual injury

Must be preheated before ignition

Normally stable but can become unstable at elevated temperatures

and pressures. 0

During a fire offers no hazard beyond

combustion

Will not burn Stable even under fire conditions.

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National Fire Protection Association (NFPA) ratings are included to indicate the types and severity of the hazards. The NFPA ratings are on a scale of 0-4 with 0 being nonhazardous and 4 being most hazardous. The ratings are described in Table 1.

Chemical Storage1 There has been much concern, and some confusion, about the proper storage of

laboratory chemicals. Here “proper” means the storage of chemicals in such a manner as to prevent incompatible materials from being accidentally mixed together in the event of the breakage of one or more containers in the storage area or to prevent the formation of reactive vapors that may require vented chemical storage areas. Below is a concise guide to the storage of common laboratory chemicals. 1) Perchloric acid is separated from all other materials. 2) Hydrofluoric acid is separated from all other materials. 3) Concentrated nitric acid is separated from all other materials.

4) Highly toxic materials (LD50 of 50 mg/kg or less) are stored separately.

5) Carcinogenic chemicals are stored separately. 6) Inorganic acids (except for 1, 2, 3 above) are stored separately. 7) Bases are stored separately. 8) Strong oxidizing agents are stored separately. 9) Strong reducing agents are stored separately. 10) Water reactive, pyrophoric and explosive materials are stored separately. 11) Flammable organic materials (solvents, organic acids, organic reagents) are stored

separately.

Guidelines for separating incompatible chemicals:

1) Place the chemicals to be stored separately in a heavy gauge Nalgene (or similar plastic) tub. Plastic secondary containers must be compatible with the material being stored.

2) Strong acids, especially perchloric, nitric and hydrofluoric are best stored in plastic containers designed to store strong mineral acids. These are available from lab equipment supply houses.

3) Bottle-in-a-can type of containers are also acceptable as secondary containment. Small containers of compatible chemicals may be stored in a dessicator or other secure container. Secondary containment is especially useful for highly toxic materials and carcinogens.

4) Dry chemicals stored in approved cabinets with doors may be grouped together by compatibility type on separate shelves or areas of shelves separated by taping off sections of shelving to designate where chemicals of one type are stored. Physically separated cabinets may be used to provide a barrier between groups of

1 Prepared by Tom Shelley (Chemical Hygiene Officer) 7/96. For additional information or questions you may have, please contact Tom Shelley at 5-4288.

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Laboratory Safety

stored incompatible chemicals. Strong mineral acids may be stored in one cabinet and strong bases stored in a second cabinet, for example. Flammable solvents should be stored in a rated flammable storage cabinet if available.

If you are uncertain of the hazardous characteristics of a particular chemical refer

to the MSDS for that material. A good MSDS will not only describe the hazardous characteristics of the chemical, it will also list incompatible materials.

Transporting Chemicals Transport all chemicals using the container-within-a-container concept to shield

chemicals from shock during any sudden change of movement. Large containers of corrosives should be transported from central storage in a chemically resistant bucket or other container designed for this purpose. Stairs must be negotiated carefully. Elevators, unless specifically indicated and so designated, should not be used for carrying chemicals. Smoking is never allowed around chemicals and apparatus in transit or in the work area itself.

When moving in the laboratory, anticipate sudden backing up or changes in direction from others. If you stumble or fall while carrying glassware or chemicals, try to project them away from yourself and others.

When a flammable liquid is withdrawn from a drum, or when a drum is filled, both the drum and the other equipment must be electrically wired to each other and to the ground in order to avoid the possible buildup of a static charge. Only small quantities should be transferred to glass containers. If transferring from a metal container to glass, the metal container should be grounded.

Chemical Disposal The Environmental Protection Agency (EPA) classifies wastes by their reaction

characteristics. A summary of the major classifications and some general treatment guidelines are listed below. Specific information may be found in the book, Prudent Practices for Disposal of Chemicals from Laboratories, as well as other reference materials.

Ignitability: These substances generally include flammable solvents and certain solids. Flammable solvents must never be poured down the drain. They should be collected for disposal in approved flammable solvent containers. In some cases it may be feasible to recover and reuse solvents by distillation. Such solvent recovery must include appropriate safety precautions and attention to potentially dangerous contamination such as that due to peroxide formation.

Corrosivity: This classification includes common acids and bases. They must be collected in waste containers that will not ultimately corrode and leak, such as plastic containers. It often may be appropriate to neutralize waste acids with waste bases and where allowed by local regulations, dispose of the neutral materials via the sanitary sewer system. Again, the nature of the neutralized material must be considered to ensure that it does not involve an environmental hazard such as chromium salts from chromic acid neutralization.

Reactivity: These substances include reactive metals such as sodium and various water reactive reagents. Compounds such as cyanides or sulfides are included in this

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class if they can readily evolve toxic gases such as hydrogen cyanide. Their collection for disposal must be carried out with particular care. When present in small quantities, it is advisable to deactivate reactive metals by careful reaction with appropriate alcohols and to deactivate certain oxygen or sulfur containing compounds through oxidation. Specific procedures should be consulted.

Toxicity: Although the EPA has specific procedures for determining toxicity, all chemicals may be toxic in certain concentrations. Appropriate procedures should be established in each laboratory for collection and disposal of these materials.

The handling of reaction byproducts, surplus and waste chemicals, and

contaminated materials is an important part of laboratory safety procedures. Each laboratory worker is responsible for ensuing that wastes are handled in a manner that minimizes personal hazard and recognizes the potential for environmental contamination.

Most instructional laboratories will have clear procedures for students to follow in order to minimize the generation of waste materials. Typically reaction byproducts and surplus chemicals will be neutralized or deactivated as part of the experimental procedure. Waste materials must be handled in specific ways as designated by federal and local regulations. University guidelines for waste disposal can be found in chapter 7 of the Chemical Hygiene Plan (available at http://www.cee.cornell.edu/safety/ )

Some general guidelines are: 1) Dispose of waste materials promptly. When disposing of chemicals one basic

principle applies: Keep each different class of chemical in a separate clearly labeled disposal container.

2) Never put chemicals into a sink or down the drain unless they are deactivated or neutralized and they are allowed by local regulation in the sanitary sewer system. [See Chemical Hygiene Plan for list of chemicals that can be safely disposed of in the sanitary sewer.]

3) Put ordinary waste paper in a wastepaper basket separate from the chemical wastes. If a piece of paper is contaminated, such as paper toweling used to clean up a spill, put the contaminated paper in the special container that is marked for this use. It must be treated as a chemical waste.

4) Broken glass belongs in its own marked waste container Broken thermometers may contain mercury in the fragments and these belong in their own special sealed "broken thermometer" container.

5) Peroxides, because of their reactivity, and the unpredictable nature of their formation in laboratory chemicals, have attracted considerable attention. The disposal of large quantities (25 g or more) of peroxides requires expert assistance. Consider each case individually for handling and disposal.

A complete list of compounds considered safe for drain disposal can be found in

Chapter 7 of the Chemical Hygiene Plan (http://www.cee.cornell.edu/safety/). Disposal techniques for chemicals not found in this list must be disposed of using techniques approved of by Cornell Environmental Health and Safety. When possible,

19

Laboratory Safety

hazardous chemicals can be neutralized and then disposed. When chemicals are produced that cannot be disposed of using the sanitary sewer, techniques to decrease the volume of the waste should be considered.

References Safety in Academic Chemistry Laboratories. A publication of the American Chemical

Society Committee on Chemical Safety. Fifth edition. 1990 Cornell University Chemical Hygiene Plan: Guide to Chemical Safety for Laboratory

Workers. A publication of the Office of Environmental Health, 2000. (http://www.ehs.cornell.edu/lrs/CHP/chp.htm)

OSHA Laboratory Standard

One of the best books to get started with regulatory compliance is a publication from the American Chemical Society entitled, "Laboratory Waste Management. A Guidebook."

Questions 1) Why are contact lenses hazardous in the laboratory? 2) What is the minimum information needed on the label for each chemical? When

are right to know labels required? 3) Why is it important to label a bottle even if it only contains distilled water? 4) Find an MSDS for sodium nitrate.

a) Who created the MSDS? b) What is the solubility of sodium nitrate in water? c) Is sodium nitrate carcinogenic? d) What is the LD50 oral rat? e) How much sodium nitrate would you have to ingest to give a 50% chance of death (estimate from available LD50 data). f) How much of a 1 M solution would you have to ingest to give a 50% chance of death? g) Are there any chronic effects of exposure to sodium nitrate?

5) You are in the laboratory preparing chemical solutions for an experiment and it is lunchtime. You decide to go to the student lounge to eat. What must you do before leaving the laboratory?

6) Where are the eyewash station, the shower, and the fire extinguishers located in the laboratory?

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Laboratory Measurements and Procedures

Introduction Measurements of masses, volumes, and preparation of chemical solutions of known

composition are essential laboratory skills. The goal of this exercise is to gain familiarity with these laboratory procedures. You will use these skills repeatedly throughout the semester.

Theory Many laboratory procedures require preparation of chemical solutions. Most

chemical solutions are prepared on the basis of mass of solute per volume of solution (grams per liter or Moles per liter). Preparation of these chemical solutions requires the ability to accurately measure both mass and volume.

Preparation of dilutions is also frequently required. Many analytical techniques require the preparation of known standards. Standards are generally prepared with concentrations similar to that of the samples being analyzed. In environmental work many of the analyses are for hazardous substances at very low concentrations (mg/L or µg/L levels). It is difficult to weigh accurately a few milligrams of a chemical with an analytical balance. Often dry chemicals are in crystalline or granular form with each crystal weighing several milligrams making it difficult to get close to the desired weight. Thus it is often easier to prepare a low concentration standard by diluting a higher concentration stock solution. For example, 100 mL of a 10 mg/L solution of NaCl could be obtained by first preparing a 1 g/L NaCl solution (100 mg in 100 mL). One mL of the 1 g/L stock solution would then be diluted to 100 mL to obtain a 10 mg/L solution.

Absorption spectroscopy is one analytical technique that can be used to measure the concentration of a compound. Solutions that are colored absorb light in the visible range. The resulting color of the solution is from the light that is transmitted. According to Beer's law the attenuation of light in a chemical solution is related to the concentration and the length of the path that the light passes through.

log oPbc

Pε =

2.1

where c is the concentration of the chemical species, b is the distance the light travels through the solution, ε is a constant Po is the intensity of the incident light, and P is the intensity of the transmitted light. Absorption, A, is defined as:

log oPA

P =

2.2

In practice Po is the intensity of light through a reference sample (such as deionized water) and thus accounts for any losses in the walls of the sample chamber. From equation 2.1 and 2.2 it may be seen that absorption is directly proportional to the concentration of the chemical species.

A bcε= 2.3

21


The instrument you will use to measure absorbance is a Hewlett Packard (HP) model 8452A diode array spectrophotometer. The diode array spectrophotometer uses a broad-spectrum source of incident light from a deuterium lamp. The light passes through the sample and is split by a grating into a spectrum of light that is measured by an array of diodes. Each diode measures a bandwidth of 2 nm with 316 diodes covering the range from 190 nm to 820 nm. The wavelengths of light and their colors are given in Table 1. The light path for the diode array spectrophotometer is shown in Figure 1.

The HP 8452A spectrophotometer has a photometric range of 0.002 - 3.3 absorbance units. In practice absorbance measurements greater than 2.5 are not very meaningful as they indicate that 99.7% of the incident light at that wavelength was absorbed. Conversely, an absorbance of 0.002 means that 0.5% of the incident light at that wavelength was absorbed.

When measuring samples of known concentration the Spectrophotometer software (page 160) calculates the relationship between absorbance and concentration at a selected wavelength. The slope (m), intercept (b) and correlation coefficient (r) are calculated using equation 2.4 through 2.6.

The slope of the best fit line is

( )2

2

x ynxy

mx

xn

∑ ∑−=

−

∑∑∑

2.4

The intercept of the line is

b my x= − 2.5

The correlation coefficient is defined as

( ) ( )2 2

2 2

x ynxy

rx y

x yn n

∑ ∑−=

− −

∑∑ ∑∑ ∑

2.6

Table 1. Wavelengths of light

color wavelength (nm) ultra violet 190-380

violet 380-450 blue 450-490

green 490-560 yellow 560-590 orange 590-630

red 630-760

Diode Array

Spectrograph Lens

ShutterGrating

SlitSample Cell

Source Lens

Deuterium Lamp

Figure 1. Diagram of light path in diode array spectrophotometer.

22


where x is the concentration of the solute (methylene blue in this exercise), y is the absorbance, and n is the number of samples.

Experimental Objectives To gain proficiency in:

1) Calibrating and using electronic balances 2) Digital pipetting 3) Preparing a solution of known concentration 4) Preparing dilutions 5) Measuring concentrations using a UV-Vis spectrophotometer

Experimental Methods

Mass Measurements Mass can be accurately measured with an electronic analytical balance. Perhaps

because balances are so easy to use it is easy to forget that they should be calibrated on a regular basis. It is recommended that balances be calibrated once a week, after the balance has been moved, or if excessive temperature variations have occurred. In order for balances to operate correctly they also need to be level. Most balances come with a bubble level and adjustable feet. Before calibrating a balance verify that the balance is level.

The environmental laboratory is equipped with balances manufactured by Denver Instruments. To calibrate the Denver Instrument balances: 1) Zero the balance by pressing the tare button. 2) Press the MENU key until "MENU #1" is displayed. 3) Press the 1 key to select Calibrate. 4) Note the preset calibration masses that can be used for calibration on the bottom

of the display. 5) Place a calibration mass on the pan (handle the calibration mass using a cotton

glove or tissue paper). 6) The balance will automatically calibrate. A short beep will occur and the display

will read CALIBRATED for three seconds, and then return to the measurement screen.

Dry chemicals can be weighed in disposable plastic "weighing boats" or other suitable containers. It is often desirable to subtract the weight of the container in which the chemical is being weighed. The weight of the chemical can be obtained either by weighing the container first and then subtracting, or by "zeroing" the balance with the container on the balance.

Temperature Measurement Use the Accumet™ pH/ion meter to measure the temperature of distilled water.

The temperature probe is the 4-mm diameter metallic probe. Place the probe in a

23


100-mL plastic beaker full of distilled water. Wait at least 15 seconds to allow the probe to equilibrate with the solution.

Pipette Technique

1) Use Figure 2 to estimate the mass of 990 µL of distilled water (at the measured temperature).

2) Use a 100-1000 µL digital pipette to transfer 990 µL of distilled water to a tared weighing boat on the 100 g scale. Record the mass of the water and compare with the expected value (Figure 2). Repeat this step if necessary until your pipetting error is less than 2%, then measure the mass of 5 replicate 990 µL pipette samples. Calculate the mean ( x defined in equation 2.7), standard deviation (s defined in equation 2.8), and coefficient of variation, s/ x , for your measurements. The coefficient of variation (c.v.) is a good measure of the precision of your technique. For this test a c.v. < 1% should be achievable.

x

xn

= ∑ 2.7

2

s

xx

nn

2( )−

=−1

∑∑ 2.8

Measure Density

1) Weigh a 100 mL volumetric flask with its cap (use the 400 g or 800 g balance).

2) Prepare 100 mL of a 1 M solution of sodium chloride in the weighed flask. Make sure to mix the solution and then verify that you have exactly 100 mL of solution. Note that the volume decreases as the salt dissolves.

3) Weigh the flask (with its cap) plus the sodium chloride solution and calculate the density of the 1 M NaCl solution.

Prepare methylene blue standards of several concentrations

1) A methylene blue stock solution of 1 g/L has been prepared. Use it to prepare 100 mL of each of the following concentrations: 1 mg/L, 2 mg/L, 3 mg/L, 4 mg/L, and 5 mg/L.

995

996

997

998

999

1000

15 20 25 30

Kg/

cubi

c m

eter

Temperature (°C) Figure 2. Density of water vs. temperature.

24


2) Note any errors in transfer of mass as you prepare these dilutions (the color will make it easy to see).

Prepare a standard curve and measure an unknown

1) See page 160 for instructions on using the UV-Vis Spectrophotometer software. 2) Measure the absorbance of the methylene blue solutions using a UV-Vis

spectrophotometer. Analyze the 5 methylene blue samples plus a distilled water sample (0 mg/L methylene blue) as standards. Select Measure Standards from the computer control palette. Fill in the information for the six samples (starting with distilled water and ending with the highest concentration of methylene blue) and follow instructions as you are prompted.

3) Save the data as \\enviro\enviro\Courses\453\fundamentals\netid_blue. 4) Record the absorbance at 660 nm for each of the solutions. You can drag the blue

cursor on the “standard graph” to the wavelength of choice and read the exact absorbance (and wavelength) in the digital display to the right of the graph. Note that you can do this after you have analyzed all of the standards.

5) Record the correlation coefficient (equation 2.6), slope (equation 2.4), and intercept (equation 2.5) for the absorbance at 660 nm vs. methylene blue concentration. These values are shown next to the “calibration graph” and correspond to the wavelength selected using the blue cursor on the “standard graph.”

6) Measure the absorbance of a methylene blue solution of unknown concentration. Select Measure Samples from the control palette. Save the data as \\enviro\enviro\Courses\453\fundamentals\netid_unknown. Record its absorbance at 660 nm and the calculated concentration. These values are given in the digital displays in the bottom left of the window.

7) Print the results by selecting Print from the control palette. 8) Export your standards spectra to the \\enviro\enviro\Courses\453\fundamentals

folder. 9) Turn on the pump and place the sipper tube in distilled water to clean out the

sample cell by selecting Run Pump from the control palette.

Prelab Questions 1) You need 100 mL of a 1 µM solution of zinc that you will use as a standard to

calibrate an atomic adsorption spectrophotometer. Find a source of zinc ions combined either with chloride or nitrate (you can use the world wide web or any other source of information). What is the molecular formula of the compound that you found? Zinc disposal down the sanitary sewer is restricted at Cornell. How does the disposal restriction for zinc influence how you prepare the zinc standard? How would you prepare this standard using techniques readily available in the environmental laboratory? Note that we have pipettes that can dispense volumes between 10 µL and 1 mL and that we have 100 mL and 1 L volumetric flasks. Include enough information so that you could prepare the standard without doing

25


any additional calculations. Consider your ability to accurately weigh small masses. Explain your procedure for any dilutions.

2) The density of sodium chloride solutions as a function of concentration is approximately 0.6985C + 998.29 (kg/m3) (C is kg of salt/m3). What is the density of a 1 M solution of sodium chloride?

Questions 1) Create a graph of absorbance at 660 nm vs concentration of methylene blue in

Excel using the exported data file. Does absorbance at 660 nm increase linearly with concentration of methylene blue?

2) Plot ε as a function of wavelength for each of the standards on a single graph. Make sure you include units and axis labels on your graph. If Beer’s law is obeyed what should the graph look like?

3) Did you use interpolation or extrapolation to get the concentration of the unknown?

4) What colors of light are most strongly absorbed by methylene blue? 5) What measurement controls the accuracy of the density measurement? What

should the accuracy be? What was your percent error in measuring the density of the 1 M NaCl solution?

27


Data Sheet

Balance Calibration

Balance ID...................................................................___________

Mass of calibration mass ..............................................___________

2nd mass used to verify calibration................................___________

Measured mass of 2nd mass.........................................___________

Temperature Measurement

Distilled water temperature...........................................___________

Pipette Technique (use DI-100 balance)

Density of water at that temperature..............................___________

Actual mass of 990 µL of pure water............................___________

Mass of 990 µL of water (rep 1) ..................................___________





Average of the 5 measurements....................................___________

Standard deviation of the 5 measurements.....................___________

Precision

Percent coefficient of variation

of the 5 measurements..................................................___________

Accuracy

average percent error for pipetting

actual mass - measured mass100

actual mass⋅ ............................___________

Where actual mass is calculated from the density of water and the setting of the pipette and measured mass is measured with the balance.

28


Measure Density (use DI-800 balance)

Molecular weight of NaCl............................................___________

Mass of NaCl in 100 mL of a 1-M solution..................

Measured mass of NaCl used.......................................___________

Measured mass of empty 100 mL flask.........................___________

Measured mass of flask + 1M solution..........................___________

Mass of 100 mL of 1 M NaCl solution.........................___________

Density of 1 M NaCl solution.......................................___________

Prepare methylene blue standards of several concentrations

Volume of 1 g/L MB diluted to 100 mL to obtain:

1 mg/L MB..................................................................___________

2 mg/L MB..................................................................___________

3 mg/L MB..................................................................___________

4 mg/L MB..................................................................___________

5 mg/L MB..................................................................___________

Measure absorbance at 660 nm using a spectrophotometer.

Spectrophotometer (computer name)?..........................___________

Absorbance of distilled water .......................................___________

Absorbance of 1 mg/L methylene blue ..........................___________





Slope at 660 nm (m) ....................................................___________

Intercept at 660 nm (b) ................................................___________

Correlation coefficient at 660 nm (r) .............................___________

Absorbance of unknown at 660 nm..............................___________

Calculated concentration of unknown............................___________

29


Lab Prep Notes

Table 2. Reagent list.

Description

Supplier

Catalog number

NaCl Fisher Scientific BP358-1 Methylene blue Fisher Scientific M291-25

Table 3. Equipment list

Description

Supplier

Catalog number

Calibra 100-1095 µL

Fisher Scientific 13-707-5

Calibra 10-109.5 µL


DI 100 analytical toploader

Fisher Scientific 01-913-1A

DI-800 Toploader Fisher Scientific 01-913-1C 100 mL volumetric Fisher Scientific 10-198-50

B UV-Vis

spectrophotometer Hewlett-Packard

Company 8452A

Table 4. Methylene Blue Stock Solution Description MW (g/M) conc. (g/L) 100 mL C16H18N3SCl 319.87 1 100.0 mg

Setup

1) Prepare stock methylene blue solution and distribute to student workstations in 15 mL vials.

2) Prepare 100 mL of unknown in concentration range of standards. Divide into two bottles (one for each spectrophotometer).

3) Verify that spectrophotometers are working (prepare a calibration curve as a test). 4) Verify that balances calibrate easily. 5) Disassemble, clean and lubricate all pipettes.

30


Reactor Characteristics

Introduction Chemical, biological and physical processes in nature and in engineered systems

usually take place in what we call "reactors." Reactors are defined by a real or imaginary boundary that physically confines the processes. Lakes, segments of a river, and settling tanks in treatment plants are examples of reactors. Most, but not all, reactors experience continuous flow (in and out). Some reactors, experience flow (input and output) only once. These are called "batch" reactors. It is important to know the mixing level and residence time in reactors, since they both affect the degree of process reaction that occurs while the fluid (usually water) and its components (often pollutants) pass through the reactor.

Tracer studies can be used to determine the hydraulic characteristics of a reactor such as the disinfection contact tanks at water treatment plants. The results from tracer studies are used to obtain accurate estimates of the effective contact time.

Reactor Classifications Mixing levels give rise to three categories of reactors; completely mixed flow

(CMF), plug flow (PF) and flow with dispersion (FD). The plug flow reactor is an idealized extreme not attainable in practice. All real reactors fall under the category of FD or CMF.

Reactor Modeling Both the CMF and the PF reactors are limiting cases of the FD reactor. Therefore the FD reactor model will be developed first. Equation 3.1 is the governing differential equation for a conservative (i.e., non-reactive) substance in a reactor that has advective transport (i.e., flow) and some mixing in the direction of flow (x - dimension).

2

d 2 -U D

C C Ct x x

∂ ∂ ∂= +

∂ ∂ ∂ 3.1

C = concentration of a conservative substance U = average fluid velocity in the x direction Dd = longitudinal dispersion coefficient t = time

The dispersion coefficient is a measure of the mixing in a system.

Flow with Dispersion One of the easiest methods to determine the mixing (dispersion) characteristics of a

reactor is to add a spike input of a conservative material and then monitor the concentration of the material in the reactor effluent.

Assuming complete mixing in y-z plane then transport occurs only in the x direction and the concentration of tracer for any x and t (after t=0) the solution to equation 3.1 gives:

31


2

C(x,t) expdd

M xD tA D tπ

′−= 44

3.2

where M = mass of conservative material in the spike, Dd = axial dispersion coefficient [L2/T], x' = x - Ut, U = longitudinal advective velocity in the reactor, and A is the cross-sectional area of the reactor. A measure of dispersion can be obtained directly from equation 3.2. From this equation we expect a maximum value of C at t =

x/U. At this time C(x,t) d

M

A D tπ=

4. If the mass of the tracer input (M) and reactor

cross-sectional area (A) are known, then Dd can be estimated. The form of equation 3.3 is exactly like the normal distribution curve:

2

2

1 exp

xx

CA xM σσ π

−= 42

3.3

where

2 2x dD tσ = 3.4

The variance in concentration over space ( 2xσ ) is the variance in concentrations

taken from many different positions in the reactor at some single moment in time, t. The variance in x ( 2

xσ ) has dimensions of length squared.

The variance of tracer concentration versus time ( 2tσ , with dimensions of time

squared) can be measured by sampling at a single point in the reactor at many different times and can be computed using the following equations.

2 2

2 20 0

0 0

t

C t t t dt t C t dtt

C t dt C t dtσ

∞ ∞

∞ ∞

( )( − ) ⋅ ( ) = = −

( ) ( )

∫ ∫

∫ ∫ 3.5

where

0

0

( )

( )

t C t dtt

C t dt

∞

∞=

∫

∫ 3.6

For discrete data points:

2

2 20

0

n

i ii

t n

ii

t C tt

C tσ =

=

⋅ ∆= −

∆

∑

∑ 3.7

and

32


0

0

n

i ii

n

ii

t C tt

C t

=

=

⋅ ∆=

∆

∑

∑ 3.8

Inlet and outlet boundary conditions affect the response obtained from a reactor. Closed reactors have little dispersion across their inlet and outlet boundaries whereas "open" reactors can have significant dispersion across their inlet and outlet boundaries. Typically open systems have no physical boundaries in the direction of flow. An example of an open system would be a river segment. Closed systems have small inlets and outlets that minimize dispersion across the inlet and outlet regions. An example of a closed system is a tank (or a lake) with a small inlet and outlet. The reactors used in the lab are closed. The t in equation 3.8 is the measured average residence time for the tracer in the reactor. For ideal closed reactors the measured residence time, t , is equal to the theoretical hydraulic residence time (θ = reactor volume/flow rate).

The above equations suggest that from the reactor response to a spike input we can compute the dispersion coefficient for the reactor. We have two options for measuring reactor response: 1) synoptic measurements: at a fixed time sampling many points along the axis of

the reactor will yield a Gaussian curve of concentration vs. distance. In practice synoptic measurements are difficult because it requires sampling devices that are time-coordinated. By combining equations 3.4, 3.7, and 3.8 it is possible to estimate the dispersion coefficient from synoptic measurements.

2) single point sampling: measure the concentration at a fixed position along the x axis of the reactor for many times. If the reactor length is fixed at L and measurements are made at the effluent of the reactor (observe the concentration of a tracer at x = L as a function of time) then x is no longer a variable and C(x,t) becomes C(t) only. The response curve obtained through single point sampling is skewed. The curve “spread” changes during the sampling period and the response curve is skewed.

Peclet Number The dimensionless parameter Pe (Peclet number) is used to characterize the level of

dispersion in a reactor. The Peclet number is the ratio of advective to dispersive transport.

d

UPe

D L= 3.9

In the limiting cases when Pe = 0 (very high dispersion) we have a complete mix regime (CMFR) and when PE = 8 (Dd = 0, no dispersion) we have a plug flow reactor (PF).

For single point sampling of the effluent response curve, skew increases as the dispersion level in the reactor increases. The degree of skew depends on the

33


dispersion coefficient, the velocity in the x-direction, and the length of the reactor. Peclet values in the range 100<Pe<8 result in a symmetric response curve.

Response curve skew makes the assumption of a symmetrical normal distribution curve inappropriate and a new relationship between the variance and the dispersion coefficient (or Pe) has to be determined. Boundary conditions affect the determination of the dispersion coefficient. The relationship between the Peclet number and variance for open systems is given by:

2 22

2 8t Pe Pe

σ θ = + ⋅

3.10

For closed systems the relationship is:

( )2 22

2 21

Pet Pe Pe eσ θ

− = − ⋅ − ⋅ 3.11

The term 2

Pein equations 3.10 and 3.11 is dominant for Peclet numbers much

greater than 10 as is shown in Figure 1. The additional terms in equations 3.10 and 3.11 are corrections for skewedness in the response curve. These skewedness corrections are not very significant for Peclet numbers greater than 10. Thus for Peclet numbers greater than 10 the Peclet number can be determined using equation 3.12 for both open and closed systems.

2

2

2

t

Peθ

σ= 3.12

Flow through Porous Media Flow through porous media (such as groundwater through soil) is a type of flow

with dispersion. The above equations can be applied by recognizing that the relevant water velocity is the pore water velocity.

The pore water velocity is U = QAε

where

A is the cross sectional area of the porous media and ε (volume of voids/total volume) is the porosity of the porous media.

Completely Mixed Flow Reactor In the case of CMF reactors, there is not

an analytical solution to the advective dispersion equation so we revert to a simple mass balance. For a completely mixed reactor a mass balance on a conservative tracer yields the following differential equation:

10000

1000

100

10

1

0.1

0.01

0.0011000010001001010.10.010.001

Pe

2/Pe

open

closed

σ2

θ 2

Figure 1. Relationship between equations 3.10 through 3.12.

34


( )in

dCV C C Q

dt= − 3.13

where Q is the volumetric flow rate and V is the volume of the reactor. Equation 3.13 can be used to predict a variety of effluent responses to tracer inputs

such as the pulse input used in this experiment. If a mass of tracer is discharged directly into a reactor so that the initial concentration of tracer in the reactor is C0 = MV

and the input concentration is zero (Cin = 0) the solution to the differential

equation is:

0

-tC C exp

t =

3.14

If a reactor has a complete mix flow regime its response (C/C0 vs. time) to a pulse input should plot as a straight line on a semi-logarithmic plot. The slope of this plot is the negative inverse of the average hydraulic residence time, t , of the reactor.

Complete mix flow regimes can be approximated quite closely in practice.

Plug Flow Reactor Plug flow regimes are impossible to attain because mass transport must be by

advection alone. There can be no differential displacement of tracer relative to the average advective velocity. In practice some mixing will occur due to molecular diffusion, turbulent dispersion, and/or fluid shear. For the case of the plug flow reactor the advective dispersion equation 3.1 reduces to:

C C

Ut x

∂ ∂= −

∂ ∂ 3.15

The velocity U serves to transform the directional concentration gradient into a temporal concentration gradient. In other words, a conservative substance moves with the advective flow of the fluid. The solutions to this differential equation for a pulse input and for a step input are shown graphically in Figure 2.

Mass Conservation When a pulse of conservative

tracer is added to a continuous flow reactor, all of the tracer is expected to leave the reactor eventually. The mass of a substance that has left the reactor is given in equation 3.16.

U

X

C

Co

U

X

C

Co

pul se i nput st ep i nput

C

Figure 2. Pulse and step input in a plug flow reactor.

35


0

n

i ii

M QC t=

= ∆∑ 3.16

where Q is the flow rate and M is the mass of any substance whose concentration is given by C. If Q and ?t are constant, then equation 3.16 can be rewritten as

0

n

ii

M Q t C=

= ∆ ∑ 3.17

Equation 3.17 can be used to determine if all of the tracer was measured in the reactor effluent.

Conductivity Measurements We will use a tracer containing salt (NaCl) and red dye # 40 (for visualization).

The concentration of NaCl will be monitored using a conductivity probe. Conductivity is the measure of a material's ability to conduct electric current. Conductivity is measured by passing an electrical current between two electrodes and then measuring the voltage. The electrodes can be made of platinum, titanium, gold-plated nickel, or graphite. Conductivity is defined as:

I

GE

= 3.18

where G is conductivity, I is the current, and E is the measured voltage. If the current is held constant, as the conductivity of the solution increases the

voltage between the electrodes will decrease. For a given current, the measured voltage will increase as the size of the electrodes decreases and as the distance between the electrodes increases. We are interested, however, in measuring properties of the solution, not properties of the conductivity probe! Specific conductivity, C, is a property of the solution.

L

C GA

= 3.19

where L is the distance between the electrodes and A is the area of the electrodes. The

term LA

is a property of the conductivity cell and is called the cell constant. In

practice, the cell constant is determined during calibration by measuring the conductivity (G) of a solution with known specific conductivity (C). The units of specific conductivity are Siemens/cm where Siemens are the inverse of Ohms.

For a solution to be conductive, it must have ions that can transport the charge between the electrodes. In pure water, the only ions available are H+ and OH-. Adding species that disassociate into charged ions increases both the concentration of ions and the specific conductivity. At low concentrations, specific conductivity increases linearly with the concentration of ions. At very high concentrations ion-ion interactions become significant and the relationship is no longer linear. The specific conductivity of several common solutions is given in Table 1.

36


Conductivity measurements are

temperature dependent. The conductivity of a solution will increase as the temperature increases. The Accumet™ meter that you will use in this laboratory compensates for this effect by also measuring the temperature and reporting the solution specific conductivity at 25°C.

In this lab sodium chloride will increase the specific conductivity of the water in the reactors. The concentration of sodium chloride will be low enough so that specific conductivity will be linearly related to the concentration of sodium chloride.

Procedures A conservative tracer will be used to characterize each of the reactors. A

conservative tracer with 20 g NaCl/L and 4 g red dye # 40/L will be used. The salt will increase the conductivity of the water and conductivity will be measured to monitor the salt concentration. The red dye was added to the tracer to make it possible to see the tracer.

A common problem when using tracers is that the tracer may have a different density than the fluid that is in the reactors. In this case the salt and dye add significantly to the density of the tracer. The tracer would tend to sink to the bottom of the reactors. To compensate for this problem the density of the water being pumped into some reactors will be increased by using a glucose solution (37 g glucose/L). Glucose is nonionic and thus will not increase the conductivity of the solution.

Calibrate Conductivity probe Calibrate the conductivity probe by placing it in a 495 mg NaCl/L standard. Press

the conductivity button on the Accumet™ meter if it is not already in the conductivity mode. Press standardize and enter 1000 µS/cm. Press enter and the meter will calibrate and return to the normal display mode.

Measure Conductivity of tracer Prepare a calibration curve for conductivity vs. concentration of the tracer

(expressed as mg/L of NaCl). The tracer has 20 g/L of NaCl. Measure the conductivity of tracer diluted with distilled water so that the final concentrations of NaCl are 500, 200, and 100 mg/L. As a zero point measure the conductivity of distilled water.

Table 1. Conductivity of some common solutions.

Solution Specific Conductivity pure water 0.055 µS/cm

distilled water 0.5 µS/cm deionized water 0.1-10 µS/cm

typical drinking water 0.5-1.0 mS/cm wastewater 0.9-9.0 mS/cm

maximum drinking water 1.5 mS/cm ocean water 53 mS/cm 10% NaOH 355 mS/cm

495 mg/L NaCl 1 mS/cm

37


Measure Reactor Response to Pulse Input For each reactor add a pulse input of sodium, measure conductivity vs. time in the

reactor effluent and use the Compumet™ software to monitor the conductivity vs. time (see discussion below). Save the collected data for later analysis using a spreadsheet program. The experimental setup is shown in Figure 3. Specific instructions for each type of reactor are detailed below.

Porous Media The porous media column is 2.5

cm in diameter, 60 cm long and contains 60 cm of glass beads. The overall porosity including headspace and underdrains is approximately 0.4. Use this information to estimate the volume of water in the reactor. The conductivity probe should be plumbed into the effluent line. 1) Verify that the flow rate is set to

10 mL/min. 2) Inject 10 mg NaCl (0.5 mL of

tracer) into the influent line. 3) Select Set Method from the

Compumet™ control palette. Use automatic data transmission with a timed interval of 1 second. Set channel A to Conductivity and channel B to Off.

4) Select Monitor Sample from the control palette. 5) Start the pump and press the enter key on the keyboard to begin data acquisition. 6) Measure the actual flow rate by collecting a timed sample from the effluent. To

get an accurate flow rate you should collect a sample for several minutes. 7) Estimate the width of the tracer pulse when the pulse nears the top of the reactor

and record the corresponding time. This information will be used to obtain an estimate of the dispersion coefficient.

8) Turn off the pump when the conductivity returns to the baseline conductivity. 9) Stop data acquisition by clicking on the Stop Sampling button. 10) Save the data to \\Enviro\enviro\Courses\453\reactors\netid_porousmedia by

selecting Save data from the control palette. The data will be saved in a file (tab delimited format) that can be opened by any spreadsheet program.

Completely Mix Flow Reactor (CMFR) 1) Verify that the flow rate is set to 300 mL/min.

Feed solution(glucosesolution)

Peristaltic pump

Flow with dispersion

or

Completely mixed reactor

Plug flow reactor

or

stirrer

Injection port

Figure 3. Reactor schematic. Only one reactor at a time will be connected to the peristaltic pump.

38


2) Fill the CMFR with distilled water to within about 2 mm of the overflow drain . 3) Measure the conductivity of the distilled water. 4) Set the stirrer to the highest setting that doesn't cause splashing (setting 8) and

place the conductivity probe near the stir bar. 5) Add 800 mg NaCl (40 mL tracer) directly to the CMFR. 6) Select Set Method from the Compumet™ control palette. Use automatic data

transmission with a timed interval of 10 second. Set channel A to Conductivity and channel B to Off.

7) Select Monitor Sample from the control palette. 8) Start the pump and press the enter key on the keyboard to begin data acquisition. 9) Record the time when water begins flowing out the overflow (this is your actual

time zero!) 10) Measure the flow rate by collecting a timed sample from the effluent. To get an

accurate flow rate you should collect a sample for several minutes. 11) Turn off the pump after 2 residence times. 12) Stop data acquisition by clicking on the Stop Sampling button. 11) Save the data to \\Enviro\enviro\Courses\453\reactors\netid_CMFR by selecting

Save data from the control palette. The data will be saved in a file (tab delimited format) that can be opened by any spreadsheet program.

13) Determine the volume of water in the CMFR.

Baffled Tank Reactor The baffled tank reactor is a simple attempt to reduce mixing and short-circuiting.

The channels are approximately 4.5 cm wide, 4.8 cm deep and have a total length of 80 cm. 1) Verify that the flow rate is set to 300 mL/min. 2) Determine the volume of water in the baffled tank. 3) Fill the baffled tank with glucose water. 4) Measure the conductivity of the glucose water. 5) Select Set Method from the Compumet™ control palette. Use automatic data


6) Select Monitor Sample from the control palette. 7) Inject 200 mg NaCl (10 mL of tracer) into the influent line with a syringe. 8) Start the pump and press the enter key on the keyboard to begin data acquisition. 9) During data acquisition, it is important to gently move the conductivity probe up

and down to continually bring the probe into contact with the changing solution. While moving the probe up and down, do not lift the probe so high that the platinum contacts leave the solution

39


10) Measure the actual flow rate by collecting a timed sample from the effluent. To get an accurate flow rate you should collect a sample for several minutes.

11) Turn off the pump when the conductivity returns to the baseline conductivity. 12) Stop data acquisition by clicking on the Stop Sampling button. 13) Measure the average conductivity of the remaining solution in the baffled tank. 12) Save the data to \\Enviro\enviro\Courses\453\reactors\netid_baffled by selecting


Pipe Flow The pipe flow setup consists of 15.24 m of 6 mm ID tubing.

1) Verify that the flow rate is set to 50 mL/min. 2) Fill the pipe with glucose water. 3) Select Set Method from the Compumet™ control palette. Use automatic data


4) Select Monitor Sample from the control palette. 5) Inject 10 mg NaCl (0.5 mL of tracer) into the influent line with a syringe. 6) Start the pump and press the enter key on the computer keyboard to begin data

acquisition. 7) Measure the actual flow rate by collecting a timed sample from the effluent. To

get an accurate flow rate you should collect a sample for several minutes. 8) Turn off the pump when the conductivity returns to the baseline conductivity. 9) Stop data acquisition by clicking on the Stop Sampling button. 13) Save the data to \\Enviro\enviro\Courses\453\reactors\netid_pipe by selecting


Prelab Questions 1) Why is a 37 g/L glucose solution used for the plug flow reactor? Why is the

glucose solution not needed for the completely mixed flow reactor? 2) Why is the conductivity of pure water not zero?

Data Analysis Use a consistent set of units throughout your data analysis and include the units in

your spreadsheet and report! 1) Derive an equation relating concentration of NaCl in the tracer to conductivity

based on the 4 point calibration curve. Use the slope from the equation and the baseline conductivity of each of the reactors to convert the conductivity data to concentration of NaCl for each reactor.

40


2) Perform a mass balance on the salt. When applicable include the measurements of residual salt left in the reactors at the end of your experiments. Use equation 3.17 to calculate the mass of NaCl measured in the effluent from each reactor and compare with the mass of NaCl added.

3) Calculate the volume of the pipe flow and porous media reactors based on their dimensions and porosity.

4) From the data determine t for each reactor (use equation 3.8 for the baffled tank, pipe flow, and porous media column and use equation 3.14 for the CMFR) and compare with the hydraulic residence time (θ = V/Q). Discuss any discrepancies.

5) The width of the plume as measured by eye for the porous media column is approximately 4 standard deviations (2 σ on each side of the peak). Use your measurement of the width of the plume and equation 3.4 to estimate the dispersion coefficient for the porous media reactor.

6) Estimate the Peclet numbers (equation 3.12) and the dispersion coefficients (equation 3.9) for the baffled tank, pipe flow, and porous media column. Compare the dispersion coefficient for the porous media reactor with the estimate obtained in the previous step.

7) Plot actual and theoretical effluent tracer concentration versus time for the three reactors. Use the calculated dispersion coefficient (equation 3.9) substituted into equation 3.2 to model the baffled tank, pipe flow, and porous media column. On the graphs also show θ, and t (these can be added to your graph in Excel as additional plots). Use the CMFR volume to obtain the theoretical value of the initial concentration.

8) Compare your results with theory. Give possible reasons for deviations from theoretical expectations.

9) Discuss what you learned.

41


Lab Prep Notes


Description Supplier Catalog number

Accumet™ 50 meter


floating stir bar Fisher Scientific 14-511-99A magnetic stirrer Fisher Scientific 11-500-7S

6 L container Fisher Scientific 03-484-22 Conductivity

Cell 1/cm Fisher Scientific 13-620-160

column Fisher Scientific K420830-6010 glass shot 297-420 µm

Sunbelt Industries

Mil 5

6 L container with baffles

CEE shop

3-port leur manifold

Cole Parmer H-06464-82

variable flow digital drive


Easy-Load pump head


PharMed tubing size 18


20 liter HDPE Jerrican

Fisher Scientific 02-961-50C

Table 3. Reagent list Description Supplier/Source Catalog

number NaCl Fisher Scientific BP358-1

red dye #40 MG Newell 07704-1 glucose Aldrich 15,896-8 1000 µS solution

495 mg NaCl/L

Tracer 20 g NaCl/L 4 g red dye #40/L

glucose feed solution

37 g glucose/L

1) Prepare glucose solution for the baffled tank and pipe flow reactors in Jerricans. 2) Prepare 1 L of tracer. 3) Prepare 1 L of 1000 µS/cm solution (conductivity standard). 4) Distribute tracer and conductivity standard for each setup. 5) Use # 18 tubing for CMFR and baffled tank. Use #16 tubing for pipe flow and

porous media reactors. 6) Place all conductivity probes in distilled water so they are conditioned. Otherwise

the readings will drift.

42


7) 0.5 mL tracer/100 mL = 100 mg/L. The concentration of glucose

required to achieve the same density as a sodium chloride solution is 1.848 times as great.

990

1000

1010

1020

1030

1040

0 20 40 60 80 100

C (g/L)

dens

ity (g

/L)

density = 0.378C + 998.215

Figure 4. Density of glucose solution as a function of glucose concentration.

995

1000

1005

1010

1015

1020

1025

0 10 20 30

C (g/L)

dens

ity (g

/L) density = 0.6985C + 998.29

Figure 5. Density of sodium chloride solution as a function of concentration.

43

Acid Precipitation and Remediation of Acid Lakes


Introduction Acid precipitation has been a serious environmental problem in many areas of the

world for the last few decades. Acid precipitation results from the combustion of fossil fuels, that produces oxides of sulfur and nitrogen that react in the earth's atmosphere to form sulfuric and nitric acid. One of the most significant impacts of acid rain is the acidification of lakes and streams. In some watersheds the soil doesn’t provides ample acid neutralizing capacity to mitigate the effect of incident acid precipitation. These susceptible regions are usually high elevation lakes, with small watersheds and shallow non-calcareous soils. The underlying bedrock of acid-sensitive lakes tends to be granite or quartz. These minerals are slow to weather and therefore have little capacity to neutralize acids. The relatively short contact time between the acid precipitation and the watershed soil system exacerbates the problem. Lakes most susceptible to acidification: 1) are located downwind, sometimes hundreds of miles downwind, from major pollution sources–electricity generation, metal refining operations, heavy industry, large population centers; 2) are surrounded by hard, insoluble bedrock with thin, sandy, infertile soil; 3) have a high runoff to infiltration ratio; 4) have a low watershed to lake surface area. Isopleths of precipitation pH are depicted in Figure 1.

Figure 1. The pH of precipitation in 1999.

44


In acid-sensitive lakes the major parameter of concern is pH (pH = -log{H+}, where {H+} is the hydrogen ion activity, and activity is approximately equal to concentration in moles/L). In a healthy lake, ecosystem pH should be in the range of 6.5 to 8.5. In most natural freshwater systems, the dominant pH buffering (controlling) system is the carbonate system. The carbonate buffering system is composed of four components: dissolved carbon dioxide ( 2 aqCO ), carbonic acid

( 2 3H CO ), bicarbonate ( -3HCO ), and carbonate ( -2

3CO ). Carbonic acid exists only at very low levels in aqueous systems and for purposes of acid neutralization is indistinguishable from dissolved carbon dioxide. Thus to simplify things we define

[ ]*2 3 2 aq 2 3H CO CO H CO = + 4.1

The 2 aqCO >> [ ]2 3H CO and thus *2 3 2 aqH CO CO ≅ (all terms enclosed in []

are in units of moles/L). The sum of all the molar concentration of the components of the carbonate system

is designated as CT as shown in equation 4.2.

* - -2T 2 3 3 3C H CO HCO CO = + + 4.2

The carbonate system can be considered to be a "volatile" system or a "non-volatile" system depending on whether or not aqueous carbon dioxide is allowed to exchange and equilibrate with atmospheric carbon dioxide. Mixing conditions and hydraulic residence time determine whether an aquatic system is volatile or non-volatile relative to atmospheric carbon dioxide equilibrium. First, consider the "non-volatile" system.

Non-volatile System For a fixed CT, the molar concentration of each species of the carbonate system is

determined by pH. Equations 4.3-4.8 show these functional relationships.

*2 3 0

1 1 22

H CO 1

[ ] [ ]

TT

CC

K K KH H

α

+ +

= = + +

4.3

where

01 1 2

2

1

1[ ] [ ]

K K KH H

α

+ +

=+ +

4.4

-3 1

2

1

HCO [ ]

1[ ]

TT

CC

KHK H

α+

+

= = + +

4.5

where

45


1α+

2+

1

1 =

Κ[Η ]+ 1 +

Κ [Η ]

4.6

-23 22

1 2 2

CO [ ] [ ]

1

TT

CC

H HK K K

α+ + = =

+ + 4.7

where

2α+ 2 +

1 2 2

1 =

[Η ] [Η ]+ +1

Κ Κ Κ

4.8

K1 and K2 are the first and second dissociation constants for carbonic acid and α0, α1, and α2 are the fraction of CT in the form *

2 3H CO , -3HCO , and -2

3CO respectively.

Because K1 and K2 are constants (K1 = 10-6.3 and K2 = 10-10.3), α0, α1, and α2 are only

functions of pH. A measure of the susceptibility of lakes to acidification is the acid neutralizing

capacity (ANC) of the lake water. In the case of the carbonate system, the ANC is exhausted when enough acid has been added to convert the carbonate species -

3HCO ,

and -23CO to *

2 3H CO . A formal definition of total acid neutralizing capacity is given by equation 4.9.

- -2 -3 3ANC HCO 2 CO OH - H+ = + + 4.9

ANC has units of equivalents per liter. The hydroxide ion concentration can be obtained from the hydrogen ion concentration and the dissociation constant for water Kw.

-OH H

wK+

= 4.10

Substituting equations 4.5, 4.7, and 4.10 into equation 4.9, we obtain

( ) HH

wT

KANC C α α +

1 2 + = + 2 + −

4.11

For the carbonate system, ANC is usually referred to as alkalinity.2

2 Alkalinity can be expressed as equivalents/L or as mg/L (ppm) of CaCO3. 50,000

mg/L CaCO3 = 1 equivalent/L.

46


Volatile Systems: Now consider the case where aqueous 2 aqCO is volatile and in equilibrium with

atmospheric carbon dioxide. Henry's Law can be used to describe the equilibrium relationship between atmospheric and dissolved carbon dioxide.

22 aq CO HCO P K = 4.12

where KH is Henry's constant for CO2 in moles/L-atm and PCO2 is partial pressure of

CO2 in the atmosphere (KH = 10-1.5 and PCO2 = 10-3.5). Because 2 aqCO is

approximately equal to *2 3H CO and from equations 4.1 and 4.3

2 0CO H TP K Cα= 4.13

2TC CO HP K

a0

= 4.14

Equation 4.14 gives the equilibrium concentration of carbonate species as a function of pH and the partial pressure of carbon dioxide.

The acid neutralizing capacity expression for a volatile system can be obtained by combining equations 4.14 and 4.11.

2 HH

CO H wP K K

ANCa

α α +1 2 +

0

= ( + 2 ) + − 4.15

In both non-volatile and volatile systems, equilibrium pH is controlled by system

ANC. Addition or depletion of any ANC component in equation 4.11 or 4.15 will result in a pH change. Natural bodies of water are most likely to approach equilibrium with the atmosphere (volatile system) if the hydraulic residence time is long and the body of water is shallow.

Lake ANC is a direct reflection of the mineral composition of the watershed. Lake watersheds with hard, insoluble minerals yield lakes with low ANC. Typically watersheds with soluble, calcareous minerals yield lakes with high ANC. ANC of freshwater lakes is generally composed of bicarbonate, carbonate, and sometimes organic matter ( -

orgA ). Organic matter derives from decaying plant matter in the watershed. When organic matter is significant, the ANC becomes (from equations 4.11 and 4.15):

-org H A

Hw

T

KANC C α α +

1 2 + = ( + 2 ) + − +

4.16

2 -orgH A

HCO H w

P K KANC

aα α +

1 2 +0

= ( + 2 ) + − + 4.17

47


where equation 4.16 is for a non-volatile system and equation 4.17 is for a volatile system.

During chemical neutralization of acid, the components of ANC associate with added acid to form protonated molecules. For example:

- *3 2 3H HCO H CO+ + → 4.18

or

-org orgH A HA+ + → 4.19

In essence, the ANC of a system is a result of the reaction of acid inputs to form associated acids from basic anions that were dissolved in the lake water. The ANC (equation 4.9) is consumed as the basic anions are converted to associated acids. This conversion is near completion at low pH (approximately pH 4.5 for the bicarbonate and carbonate components of ANC). Neutralizing capacity to another (probably higher) pH may be more useful for natural aquatic systems. Determination of ANC to a particular pH is fundamentally easy — simply add and measure the amount of acid required to lower the sample pH from its initial value to the pH of interest. Techniques to measure ANC are described under the procedures section of this lab.

Neutralization of acid precipitation can occur in the watershed or directly in the lake. How much neutralization occurs in the watershed versus the lake is a function of the watershed to lake surface area. Generally, watershed neutralization is dominant. Recently engineered remediation of acid lakes has been accomplished by adding bases such as limestone, lime, or sodium bicarbonate to the watershed or directly to the lakes.

Experimental Objectives

Phase I: Acid Neutralization by Soil Column In this experiment, we will study the effect of watershed soil on neutralization of

acid precipitation. Simulated acid precipitation (dilute sulfuric acid) will be applied to a column of watershed soil that will discharge to a volume of water that represents a lake. The column will contain an artificial soil amended with CaCO3 representing a highly calcareous soil. Influent and effluent lake pH and ANC will be measured over time to document the depletion of the ANC of the soil column and the subsequent acidification of the lake. Organic matter as an ANC component will not be incorporated into these experiments.

Phase II. Acid Lake Remediation Remediation of acid lakes involves addition of ANC so that the pH is raised to an

acceptable level and maintained at or above this level for some design period. In this experiment sodium bicarbonate (NaHCO3) will be used as the ANC supplement. Since ANC addition usually occurs as a batch addition, the design pH is initially exceeded. ANC dosage is selected so that at the end of the design period pH is at the acceptable level. Care must be taken to avoid excessive initial pH — high pH can be as deleterious as low pH.

48


The most common remediation procedure is to apply the neutralizing agent directly to the lake surface, instead of on the watershed. We will follow that practice in this lab experiment. Sodium bicarbonate will be added directly to the surface of the lake that has an initial ANC of 50 µeq/L and is receiving acid rain with a pH of 3. After the sodium bicarbonate is applied, the lake pH and ANC will be monitored for approximately one hour.

Experimental Apparatus The experimental apparatus consists of an acid rain storage reservoir, peristaltic

pump, soil column, and lake (Figure 2). The pH of the lake influent and of the lake will be monitored using pH probes connected to a pH meter. The soil column will be removed for phase II of the experiment.

The soil column is filled with glass beads. The ANC of the soil column is the result of adding CaCO3 slurry to the top of the soil column. Undissolved CaCO3

quickly settles to the glass beads and thus CaCO3 is only removed from the column as it dissolves. For this experiment, 250 mg of CaCO3 has been added to the soil column. Detail of the soil column is shown in Figure 3.

Experimental Procedures

Calibration of pH Meter Calibrate both pH probes

attached to the pH meter. To calibrate the probe attached to channel A toggle the display by pressing Channel until only the pH from channel A is in the display. (The display toggles between channel A, channel B, and both channels.) Then follow the calibration procedure outlined in the Appendix of this manual (page 157). Repeat the procedure for the channel B probe (if you are doing the soil column experiment).

Feed solution

Peristaltic pump

Soil column

Lake

Lake effluent Figure 2. Schematic drawing of the experimental setup.

10 cm IDPlexiglas tube

PVC cap

PVC grid

Adaptor

Wire cloth Wire screen

Glassbeads

Underdrain

Plexiglas baffle

1/8 in 'O' ring

Stainless steel fastener attachingPlexiglas cover to the filter base

Plexiglas cover

6 mm diameterPlexiglas dowel

Figure 3. Soil column used for neutralizing acid rain.

49


Acid Neutralization by Soil Column Experiment In this experiment, the "acid rain" will pass through the soil column before running

into the lake. Flow should be controlled to give a 15 minutes hydraulic retention time in the lake. With a lake volume of 5 liters, the required flow rate is 334 mL/min. Since surface waters are seldom at equilibrium or steady state, this experiment will be run under dynamic conditions. The lake will start out with no ANC. Data will be taken from the onset of flow. 1) Measure the volume of the lake (The lake weighs too much for the 5 kg balance

but it can be weighed in two parts. Weigh the empty lake container, fill the lake, pour half of the lake into a weighed container, weigh both the lake container and the second container, add the weights of the containers with water and subtract the container weights.)

2) Fill lake with distilled water. 3) Add 1 mL of bromocresol green indicator to the lake. 4) Preset pump to give desired flow rate of 334 mL/min (5 L/15 minutes). 5) Measure the pH of the acid rain. 6) Prepare to monitor the pH of two probes using the Compumet™ software. Set the

method for automatic sampling of pH probes on both channel A and channel B every 10 seconds. See section on "Compumet™ software" (page 157) for information on using the Computer™ software to monitor pH.

7) Place the probe attached to channel A to monitor influent to the lake. Place the probe attached to channel B to monitor the pH of the lake.

8) Label sample bottles (see step 13). 9) Set stirrer speed to setting 8.

10) Observe the CaCO3 on top of the soil column.

11) At time equal zero start the pump and begin monitoring pH at the lake influent and in the lake.

12) Measure the flow rate using 100-mL volumetric flask and a stopwatch. 13) Take 50-mL grab samples from the lake influent and effluent at 5, 10, 15, and 20

minutes. The sample volumes do not need to be measured. Collect the samples in 125-mL bottles.

14) Observe the depletion of the CaCO3 on top of the soil column.

15) After the 20-minute sample turn off the pump and stop sampling pH. 14) Save the pH data to \\Enviro\enviro\Courses\453\acid\netid_column by selecting


Acid Lake Remediation Experiment In this experiment sodium bicarbonate will be added to a lake to mitigate the

deleterious effect of acid rain. Usually sodium bicarbonate is added in batch doses (as opposed to metering in). The quantity of sodium bicarbonate added depends on how

50


long a treatment is desired, the acceptable pH range and the quantity and pH of the incident rainfall. For purposes of this experiment, a 15-minute design period will be used. That is, we would like to add enough sodium bicarbonate to keep the lake at or above its original pH and alkalinity for a period of 15 minutes (i.e. for one hydraulic residence time).

By dealing with ANC instead of pH as a design parameter, we avoid the issue of whether the system is at equilibrium with atmospheric carbon dioxide. Keep in mind that eventually the lake will come to equilibrium with the atmosphere. In practice, neutralizing agent dosages may have to be adjusted to take into account non-equilibrium conditions.

We must add enough sodium bicarbonate to equal the negative ANC from the acid precipitation input plus the amount of ANC lost by outflow from the lake during the 15-minute design period. Initially (following the dosing of sodium bicarbonate) the pH and ANC will rise, but over the course of 15 minutes, both parameters will drop. Calculation of required sodium bicarbonate dosage requires performing a mass balance on ANC around the lake. This mass balance will assume a completely mixed lake and conservation of ANC. Chemical equilibrium can also be assumed so that the sodium bicarbonate is assumed to react immediately with the incoming acid precipitation.

( )in out

d(ANC)Q ANC - ANC V

dt= 4.20

where: ANCout = ANC in lake outflow at any time t (for a completely mixed lake the

effluent ANC is the same as the ANC in the lake) ANCin = ANC of acid rain input V = volume of reactor Q = acid rain input flow rate.

If the initial ANC in the lake is designated as ANC0, then the solution to the mass balance differential equation is:

( ) 01-t/ -t/

out inANC ANC - ANCe eθ θ

= ⋅ + ⋅ 4.21

where: θ = V/Q We want to find ANC0 such that ANCout = 50 µeq/L when t is equal to θ. Solving

for ANC0 we get

( )0 out inANC ANC - ANC 1 -t/ t/ - e eθ θ = ⋅

4.22

The ANC of the acid rain (ANCin) can be estimated from its pH. Below pH 6.3 most of the carbonates will be in the form *

2 3H CO and thus for pH below about 4.3 equation 4.9 simplifies to

ANC H+ ≅ − 4.23

51


An influent pH of 3.0 implies the ANCin = - H+ = -0.001

Substituting into equation 4.22:

( )1 10ANC 0.000050 0.001 1 - - e e = + ⋅

= 1.854 meq/L

The quantity of sodium bicarbonate required can be calculated from:

[NaHCO3]0 =ANC0 4.24

where [NaHCO3]0 = moles of sodium bicarbonate required per liter of lake water

3 33

3

1.854 mmole NaHCO 84 mg NaHCO× × 5 Liters = 779 mg NaHCO

liter mmole NaHCO 4.25

1) Verify that the system is plumbed so that the “acid rain” is pumped directly into the lake.

2) Take a 50-mL sample from the acid rain container. Collect the sample in a 125-mL bottle.

3) Preset pump to give desired flow rate of 334 mL/min (5 L/15 minutes). 4) Fill lake with distilled water. 5) Set stirrer speed to 8. 6) Add 1 mL of bromocresol green indicator solution to the lake.

7) Weigh out 779 mg (not grams!) NaHCO3.

8) Add NaHCO3 to the lake.

9) After the lake is well stirred take a 100 mL sample from the lake. 10) Prepare to monitor the pH of one probe using the Compumet™ software. Set the

method for automatic sampling of the pH probe on channel A every 10 seconds. See section on "Compumet™ software" (page 157) for information on using the computer to monitor pH.

11) Place the probe attached to channel A to monitor the pH of the lake. 12) Label sample bottles (see step 13). 13) At time equal zero start the pump and begin monitoring the lake pH. 14) Take 100-mL grab samples from the lake effluent at 5, 10, 15, and 20 minutes.

The sample volumes do not need to be measured. Collect the samples in 125-mL bottles.

15) Measure the flow rate. 16) After the 20-minute sample turn off the pump and stop sampling pH. 17) Save the pH data to \\Enviro\enviro\Courses\453\acid\netid_remediate by

selecting Save data from the control palette. The data will be saved in a file (tab delimited format) that can be opened by any spreadsheet program.

18) Measure the lake volume.

52


Analytical Procedures pH. pH (-log{H+}) is usually measured electrometrically with a pH meter. The pH

meter is a null-point potentiometer that measures the potential difference between an indicator electrode and a reference electrode. The two electrodes commonly used for pH measurement are the glass electrode and a reference electrode. The glass electrode is an indicator electrode that develops a potential across a glass membrane as a function of the activity ,~ molarity, of H+. Combination pH electrodes, in which the H+-sensitive and reference electrodes are combined within a single electrode body will be used in this lab. The reference electrode portion of a combination pH electrode is a [Ag/AgCl/4M KCl] reference. The response (output voltage) of the electrode follows a "Nernstian" behavior with respect to H+ ion activity.

0

0 lnHRT

E EnF H

+

+

= +

4.26

where R is the universal gas constant, T is temperature in Kelvin, n is the charge of the hydrogen ion, and F is the Faraday constant. E0 is the calibration potential (Volts), and E is the potential (Volts) measured by the pH meter between glass and reference electrode. The slope of the response curve is dependent on the temperature of the sample and this effect is normally accounted for with simultaneous temperature measurements.

The electrical potential that is developed between the glass electrode and the reference electrode needs to be correlated with the actual pH of the sample. The pH meter is calibrated with a series of buffer solutions whose pH values encompass the range of intended use. The pH meter is used to adjust the response of the electrode system to ensure a Nernstian response is achieved over the range of the calibration standards. Refer to the appendix (page 157) for specific procedures.

To measure pH the electrode(s) are submersed in at least 50 mL of a sample. Samples are generally stirred during pH reading to establish homogeneity, to prevent local accumulation of reference electrode filling solution at the interface near the electrode, and to ensure the diffusive boundary layer thickness at the electrode surface is uniform and small.

ANC. The most common method to determine ANC for aqueous samples is titration with a strong acid to an endpoint pH. A pH meter is usually used to determine the endpoint or "equivalence point" of an ANC titration. Determination of the endpoint pH is difficult because it is dependent on the magnitude the sample ANC. Theoretically this endpoint pH should be the pH where all of the ANC of the system is consumed, but since the ANC is not known a-priori, a true endpoint cannot be predetermined. However, if most of the ANC is composed of carbonate and bicarbonate this endpoint is approximately pH = 4.5 for a wide range of ANC values.

A 100-mL sample is usually titrated under slowly stirred conditions. Stirring is accomplished by a magnetic stirrer. pH electrodes are ordinarily used to record pH as a function of the volume of strong acid titrant added. The volume of strong acid required to reach the ANC endpoint (pH 4.5) is called the "equivalent volume" and is used in the following equation to compute ANC.

53


?(equivalent vol.)(normality of titrant)

ANC =(vol. of sample)

4.27

A more accurate technique to measure ANC is the Gran plot analysis. This is the subject of a subsequent experiment. All ANC samples should be labeled and stored for subsequent Gran analysis.

Prelab Questions 1) How many grams of NaHCO3 would be required to keep the ANC levels in a lake

above 50 µeq/L for 3 hydraulic residence times given an influent pH of 3.0 a lake volume of 5 L, if the current lake ANC is 50 µeq/L?

Data Analysis K1 = 10-6.3, K2 = 10-10.3, KH = 10-1.5 mol/atm L, PCO2

= 10-3.5 atm, and Kw = 10-14.

Soil Column Experiment

1) Plot pH of both lake and soil column effluent versus time. 2) How long did it take for the ANC of the soil column to be depleted? (At a pH of

4.5 the ANC is approximately zero.) 3) How long did it take for the ANC of the lake to be depleted?

Acid Lake Remediation Experiment

1) Plot measured pH of the lake versus time. 2) Given that ANC is a conservative parameter and that the lake is essentially in a

completely mixed flow regime equation 4.21 applies. Graph the predicted ANC based on the completely mixed flow reactor equation with the plot labeled (in the legend) as “conservative ANC”.

3) Derive an equation for CT (the concentration of carbonate species) as a function of time based on the input of NaHCO3 and its dilution in the completely mixed lake assuming a non-volatile system (the equation will be the same form as equation 4.21).

4) Combine your equation for CT with equation 4.11 and plot the predicted ANC of the lake vs. time for a non-volatile system based on the measured pH (plot on the same graph as #2) with plot labeled as “non-volatile model”).

5) Plot the predicted ANC of the lake vs. time for a volatile system using equation 4.15 based on the measured pH (plot on the same graph as #2) with plot labeled as “volatile model”).

6) Compare the plots and determine whether the lake is best modeled as a volatile or non-volatile system. What changes could be made to the lake to bring the lake into equilibrium with atmospheric CO2?

54


Questions 1) What do you think would happen if enough NaHCO3 were added to the lake to

maintain an ANC greater than 50 µeq/L for 3 residence times with the stirrer turned off?

2) What are some of the complicating factors you might find in attempting to remediate a lake using CaCO3? Below is a list of issues to consider. • extent of mixing • solubility of CaCO3 (find the solubility and compare with NaHCO3) • density of CaCO3 slurry (find the density of CaCO3)

References Driscoll, C.T., Jr. and Bisogni, J.J., Jr., "Weak Acid/Base Systems in Dilute Acidified

Lakes and Streams of the Adirondack Region of New York State," in Modeling of Total Acid Precipitation Impacts J.L. Schnoor (ed.), Butterworth, Stoneham, MA., 53-72 (1983).

Driscoll, C.T., Baker, J.P., Bisogni, J.J., And Schofield, C.L., "Aluminum Speciation and Equilibria in Dilute Surface Waters of the Adirondack Region of New York State," in Geological Aspects of Acid Deposition O.P. Bricker (ed.), Butterworth, Stoneham, MA., 55-75 (1984).

Barnard. T.E., And Bisogni, J.J., Jr., "Errors in Gran Function Analysis of Titration Data for Dilute Acidified Water," Water Research, 19, No. 3 393-399 (1985).

Bisogni, J.J., Jr. and Barnard, T.E., "Numerical Technique to Correct for Weak Acid Errors in Gran Function Analysis of Titration Data," Water Research, 21, No. 10, 1207-1216 (1987).

Bisogni, J.J., Jr., "Fate of Added Alkalinity During Neutralization of an Acid Lake," Journal Environmental Engineering, ASCE, 114, No. 5, 1219-1224 (1988).

Bisogni, J.J., Jr., and Kishbaugh, S.A., "Alkalinity Destruction by Sediment Organic Matter Dissolution During Neutralization of Acidified Lakes," Water, Air and Soil Pollution, 39, 85-95 (1988).

Bisogni, J.J., Jr. and Arroyo, S.L., "The Effect of Carbon Dioxide Equilibrium on pH in Dilute Lakes," Water Research, 25, No. 2, 185-190 (1991).

Olem, H. Liming Acidic Surface Waters. Lewis Publishers, Chelsea, MI. (1991). Stumm, W. and Morgan, J.J., Aquatic Chemistry, John Wiley & Sons, Inc. NY, NY

1981.

55


Lab Prep Notes

Bromocresol Green Indicating Solution

Prepare solution of 400 mg Bromocresol green/100 mL ethanol. Add 0.2 mL of indicator solution per liter of acid rain or lake.

Acid rain Acid rain is at pH 3.0. Prepare

from distilled water. Add 1 meq H2SO4/L ([H+] at pH 3.0) to obtain a pH of 3.0. To acidify 20 liters of distilled water using 5 N H2SO4:

2 4

2 42 4

1 meq HSO 1 1N L20 L = 4 mL of 5 N HSO

L 5 N HSO 1000 meq⋅ ⋅ ⋅

Add 4 mL of bromocresol green

indicating solution to 20 L of acid rain solution.

Flow Rate The residence time of the lake

should be 15 minutes. The lake volume is 5 L. thus the flow rate is 334 mL/min. Use # 18 PharMed tubing.

Neutralizing Column Neutralization of the acid rain will

be accomplished in a porous media column amended with CaCO3. The porous media will be glass beads or sand (approximately 200-µm diameter) with no ANC. If new glass beads are used, they should be washed with distilled water to remove ANC. To neutralize the acid rain 1 meq/L of CaCO3 will be added to the column. This will be enough ANC to raise the pH to near neutral. To neutralize 5 liters (15 minutes):

3 32 43

2 4 3

1 meq CaCO 50 mg CaCO1 meq H SO5 L 250 mg CaCO

L 1meq H SO meq CaCO⋅ ⋅ ⋅ =

Table 1. Reagents


HCL 5.0 N Fisher Scientific LC15360-2 H2SO4 5N Fisher Scientific LC25840-2

CaCO3 Fisher Scientific C63-3 Na2CO3 Fisher Scientific S263-500

Buffer-Pac Fisher Scientific SB105 NaHCO3 Fisher Scientific S233-500

Bromocresol Green

Fisher Scientific B383-5

ethanol Fisher Scientific A962P-4


Description Supplier Catalog

number magnetic stirrer Fisher Scientific 11-500-7S floating stir bar Fisher Scientific 14-511-99A Accumet™ 50

pH meter Fisher Scientific 13-635-50

100-1095 µL pipette


10-109.5 µL pipette


pH electrode Fisher Scientific 13-620-108 6 L container

(lake) Fisher Scientific 03-484-22

Easy load pump head


digital pump drive

Cole Parmer H-07523-30_



soil column see design schematic,

Figure 3



56


The porous media column amended with CaCO3 will be prepared as follows.

1) Add 6 cm of glass beads (approximate) to 10 cm diameter column. 2) Fill from bottom of column with distilled water to 1 cm above glass bead surface. 3) Attach cover and fill column to top with distilled water.

4) Prepare CaCO3 slurry (approximately 25 mL distilled water + 250 mg CaCO3).

5) Pour slurry into 50 mL syringe. 6) Connect syringe to manifold at influent of column. 7) Inject slurry into column 15 minutes before use. If prepared too long in advance

the CaCO3 washes out of the column.

8) Allow CaCO3 to settle for 10 minutes before beginning to use.

Setup

1) Prepare 20-L acid rain for each group. 2) Prepare neutralizing columns. 3) Prepare bromocresol green solution if necessary. 4) Attach one Easy-Load pump head to the pump drives and plumb with #18 tubing. 5) Plumb Jerrican to pump to soil column to lake using quick connectors (see Figure

2). 6) Verify that pH probes are operational, stable, and can be calibrated. If you are

doing the soil column experiment you need 2 probes per group. 7) Verify that buffers (pH = 4, 7, 10) are distributed to each student group. 8) Provide a mount for the pH probe in the lake. 9) Set up some lakes with aeration!

57

Measurement of Acid Neutralizing Capacity


Introduction Acid neutralizing capacity (ANC) is a measure of the ability of water to neutralize

acid inputs. Lakes with high ANC (such as Cayuga Lake) can maintain a neutral pH even with some acid rain input whereas lakes with an ANC less than the acid input will not maintain a neutral pH. In the Adirondack region of New York State, lakes typically receive large inputs of acids during the spring thaw when the accumulated winter snow melts and runs off into the lakes. The ANC of Adirondack lakes is not always sufficient to neutralize these inputs.

Theory The ANC for a typical carbonate-containing sample is defined as:

- -2 -3 3ANC [HCO ] 2[CO ] [OH ] - [H ]+= + + 5.1

This equation can be derived from a charge balance if ANC is considered to be the cation contributed by a strong base titrant and if other ions present do not contribute significantly.

Determination of ANC or Alkalinity involves determination of an equivalence point. The equivalence point is defined as the point in the titration where titrant volume that has been added equals the "equivalent" volume (Ve). The equivalent volume is defined as:

·

= s se

t

V NV

N 5.2

where: Ns = normality (in this case Alkalinity or ANC) of sample, equivalents/L Vs = volume of sample, liters Nt = normality of titrant, equivalents/L.

The titration procedure involves incrementally adding known volumes of

standardized normality strong acid (or base) to a known volume of unknown normality base (or acid). When enough acid (or base) has been added to equal the amount of base (or acid) in the unknown solution we are at the "equivalence" point. (Note: the point at which we add exactly an equivalent or stoichiometric amount of titrant is the equivalence point. Experimentally, the point at which we estimate to be the equivalence point is called the titration endpoint).

There are several methods for determining Ve (or the equivalence point pH) from titration data (titrant volume versus pH). The shape of the titration curve (Vt versus pH) can reveal Ve. It can be shown that one inflection point occurs at t eV V= . In the case of monoprotic acids, there is only one inflection in the pH range of interest. Therefore, an effective method to find the equivalence volume is to plot the titration curve and find the inflection point. Alternately, plot the first derivative of the titration plot and look for a maximum.

58


Gran Plot Another method to find the ANC of an unknown solution is the Gran plot

technique. When an ANC determination is being made, titration with a strong acid is used to "cancel" the initial ANC so that at the equivalence point the sample ANC is zero. The Gran plot technique is based on the fact that further titration will result in an increase in the number of moles of H+ equal to the number of moles of H+ added. Thus after the equivalence point has been reach the number of moles of H+ added equals the number of moles of H+ in solution.

( ) ( )t t e s tN V V V V H + − = + 5.3

Solving for the hydrogen ion concentration:

( )

( )t t e

s t

N V VH

V V+ −

= + 5.4

Equation 5.4 can be solved directly for the equivalent volume.

( )s t

e tt

H V VV V

N

+ + = − 5.5

Equation 5.5 is valid if enough titrant has been added to neutralize the ANC. A better measure of the equivalent volume can be obtained by rearranging equation 5.4 so that linear regression on multiple titrant volume - pH data pairs can be used.

( )s t t t t e

s s s

V V N V N VH

V V V++

= − 5.6

We define F1 (First Gran function) as:

1F [H ]s t

s

V VV

++= 5.7

If F1 is plotted as a function of Vt the result is a straight line with slope =

t

s

NV

and abscissa intercept of Ve

(Figure 1). The ANC is readily obtained

given the equivalent volume. At the equivalence pt:

s e tV ANC V N= 5.8

Equation 5.8 can be rearranged to obtain ANC as a function of the equivalent volume.

y = 9.57E-04x - 4.62E-03R2 = 9.99E-01

00.00010.00020.00030.00040.00050.00060.00070.00080.0009

0 1 2 3 4 5 6

Volume of Titrant (mL)

Gran Function

Gran Function Linear Region

Figure 1. Gran plot from titration of a weak base with 0.05 N acid. Ct = 0.001 moles of carbonate and sample volume is 48 mL. The equivalent volume is 4.8 mL. From equation 5.9 the ANC is 5 meq/L.

59


ANC e t

s

V NV

= 5.9

pH Measurements The pH can be measured either as activity ({H+} as measured approximately by pH

meter) or molar concentration ([H+]). The choice only affects the slope of F1 since [H+] = {H+}/γ.

1{ }

F [H ] s t s t t et

s s s

V V V V V VHV V Vγ

+++ + −

= = = Ν 5.10

where γ is the activity correction factor and the slope is Nt/V0. If H+ concentration is used then

1 tF {H } Ns t t e

s s

V V V VV V

γ++ −= = 5.11

where the slope is t

sVγ ⋅ Ν

.

(This analysis assumes that the activity correction factor doesn't change appreciably during the titration).

There are many other Gran functions that can be derived. For example, one can be derived for Acidity or the concentration of a single weak or strong acid or base.

To facilitate data generation and subsequent Gran plot construction and analysis pH versus titrant volume can be read directly into a computer, that can be programmed to analyze the data using the Gran, plot theory. The program generates the Gran function for all data and then systematically eliminates data until the Gran function (plot) is as linear as possible. The line is then extrapolated to the abscissa to find the equivalent volume.

ANC Determination for Samples with pH < 4 After the equivalence point has been reached (adding more acid than ANC = 0) the

only significant terms in equation 5.1 are H+ and ANC.

- -2 -3 3H HCO 2 CO OH + >> + + 5.12

When the pH is 2 pH units or more below the pKs of the bases in the system the only species contributing significantly to ANC is the hydrogen ion (equation 5.12) and thus the ANC is simply

ANC - [H ]+= 5.13

For a sample containing only carbonates, if the pH is below 4 the ANC is approximately equal to -[H+] and no titration is necessary.

60


Titration Techniques Operationally, the first few titrant volumes can be relatively large increments since

the important data lies at pH values less than that of the equivalence point (approximately pH = 4.5 for an Alkalinity titration). As the pH is lowered by addition of acid the ionic strength of the solution increases and the activity of the hydrogen ion deviates from the hydrogen ion concentration This effect is significant below pH 3 and thus the effective linear range is generally between pH 4.5 and pH 3.0. The maximum incremental titrant volume (?Va) that will yield n points in this linear region is obtained as follows. If Vs » Vt then equation 5.3 reduces to

t

( )N [H ]t e

s

V VV

+−≅ 5.14

Let [H+]e be the concentration of hydrogen ions at the equivalence point and [H+]f be the final concentration of hydrogen ions at the end of the titration.

t e f

( ) ( )N [H ] - [H ]e e f e

s

V V V V

V+ +− − −

= 5.15

Thus the volume of acid added to go from [H+]e to [H+]f is

( )

f e

[ ] [ ]V - V

s f e

t

V H H

N

+ +−= 5.16

To obtain n data points between [H+]e - [H+]f requires the incremental titrant volume (?Vt) be 1/n times the volume of acid added between the equivalence point and the final titrant volume. Thus by substituting n?Vt, and typical hydrogen ion concentrations of [H+]e = 10-4.5 and [H+]f = 10-3.0 into equation 5.16 the maximum incremental titrant volume is obtained.

t

(0.001 0.00003) 0.001V

s s

t t

V Vn N n N

−∆ ≅ ≅ 5.17

Procedure

Calibrate the pH Meter Calibrate the pH meter using a pH probe connected to channel A. Use 3 standards

(pH = 4, 7, and 10).

Determine ANC of a Known Standard

1) Weigh a 100 mL plastic beaker.

2) Add approximately 50 mL of a 2.5 mM solution of Na2CO3 to the beaker.

3) Weigh the beaker again to determine the exact volume of Na2CO3 solution.

61


4) Place the beaker on the magnetic stirrer, add a stir bar and stir slowly. 5) Place both the pH electrode and the temperature probe in the Na2CO3 solution

using the probe holding arm attached to the Accumet™ meter. 6) Analyze the sample using Gran plot analysis as detailed at

http://www.cee.cornell.edu/mws/Software/Compumet.htm) Add 0.05 N HCl (the titrant) using a digital pipette in increments of 0.25 mL.

15) Save the pH data to \\Enviro\enviro\Courses\453\acid\netid_gran by selecting Save data from the control palette. The data will be saved in a file (tab delimited format) that can be opened by any spreadsheet program. You will use this data to plot a titration curve and to verify that the Gran technique accurately measures the ANC of a sample.

7) Record the ANC and the equivalent volume.

Determine ANC of Acid Rain Samples Determine ANC for all samples collected from the previous week's lab. Use the

same technique as outlined above (Determine ANC of known standard) except substitute the samples collected last week and use titrant increment of 0.1 mL in the linear region. For samples that have a high ANC you can reduce the analysis time by adding titrant in larger volumes initially until the pH approaches 5. If the initial pH is less than 4.5 no titration is necessary and equation 5.13 can be used to calculate the ANC.

Record the initial pH (prior to adding any titrant) and initial sample volume. After the Gran plot analysis record the alkalinity (ANC) and equivalent volume for each sample. There is no need to save the data to disk.

Prelab Questions 1) Compare the ability of Cayuga lake and Wolf pond (an Adirondack lake) to

withstand an acid rain runoff event (from snow melt) that results in 20% of the original lake water being replaced by acid rain. The acid rain has a pH of 3.5 and is in equilibrium with the atmosphere. The ANC of Cayuga lake is 1.6 meq/L and the ANC of Wolf Pond is 70 µeq/L. Assume that carbonate species are the primary component of ANC in both lakes, and that they are in equilibrium with the atmosphere. What is the pH of both bodies of water after the acid rain input? Remember that ANC is the conservative parameter (not pH!).

2) What is the ANC of a water sample containing only carbonates and a strong acid that is at pH 3.2?

3) Why is [H+] not a conserved species?

Questions 1) Plot the titration curve of 2.5 mM Na2CO3 with 0.05 N HCl (plot pH as a function

of titrant volume). Label the equivalent volume of titrant. Label the 2 regions of the graph where pH changes slowly with the dominant reaction that is occurring. Note that in a third region of slow pH change no significant reactions are occurring (added hydrogen ions contribute directly to change in pH).

62


2) Prepare a Gran plot using the data from the titration curve of the 2.5 mM Na2CO3. Use linear regression on the linear region or simply draw a straight line through the linear region of the curve to identify the equivalent volume. Compare your calculation of Ve with that calculated by the Compumet™ computer program.

3) Compare the measured ANC with the theoretical value for the 2.5 mM Na2CO3 solution. Note that ANC can be defined as the excess of positive charges over the anions of strong acids.

4) Plot the ANC of the influent and the lake from phase I of the previous lab. 5) Plot the ANC of the lake from phase II of the previous lab on the same graph as

was used to plot the conservative ANC model (see questions 2) to 5) on page 53). Did the measured ANC values agree with the conservative ANC model?

References Sawyer, C.N., P.L. McCarty and G.F. Parkin Chemistry for Environmental

Engineering, 4th ed., McGraw-Hill (1994). Pankow, J.F. Aquatic Chemistry Concepts, Lewis Publishers (1991). Morel, F.M.M. and J.G. Hering Principles and Applications of Aquatic Chemistry

Wiley-Interscience (1993). Stumm, W. and J.J. Morgan Aquatic Chemistry 2nd ed. Wiley Interscience (1981).

63


Lab Prep Notes



HCl 5.0 N Fisher Scientific LC15360-2 Buffer-Pac Fisher Scientific SB105

Na2CO3 Fisher Scientific BP357-1

Table 2. Equipment list Description Supplier Catalog

number Accumet™ 50


pH electrode Fisher Scientific 13-620-108 7x7 stirrer Fisher Scientific 11-500-7S

stirbar 1/2" long Fisher Scientific 14-511-62 100 mL Fisher

beaker Fisher Scientific 02-593-50B

Setup

1) Prepare 1 L of the known standard (2.5 mM solution of Na2CO3). The MW is 105.99 g/mole. 2.5mM

L·

105.99mgmM

= 265 mg Na2CO3/L

2) Prepare 1 L of the titrant (0.05 N HCl from 5.0 N HCl). Dilute 10 mL of 5.0 N HCl to 1 L. Distribute 100 mL titrant to each student group.

3) Verify that the pH probes are operational, stable, and can be calibrated. 4) Verify that buffers (pH = 4, 7, 10) are distributed to each student group

64


Phosphorus Determination using the Colorimetric Ascorbic Acid Technique

Introduction Phosphorus has been identified as a prime nutrient needed for algae growth in

inland environments. In 1992, the EPA reported that accelerated eutrophication was one of the leading problems facing the Nation's lakes and reservoirs. Eutrophication caused by the overabundance of nutrients in water can result in a variety of water-quality problems, including fish kills, noxious tastes and odors, clogged pipelines, and restricted recreation. In freshwater, phosphorus is often the nutrient responsible for accelerated eutrophication. Many algae blooms in rivers and lakes are attributed to elevated phosphorus concentrations resulting from human activities. Phosphorus enters surface waters from agricultural and urban runoff as well as from industrial and municipal wastewater treatment plant effluent.

No national criteria have been established for concentrations of phosphorus compounds in water; however, to control eutrophication, the EPA makes the following recommendations:

• Total phosphates should not exceed 50 µg/L (as phosphorus) in a stream at a point where it enters a lake or reservoir.

• Total phosphorus should not exceed 100 µg/L in streams that do not discharge directly into lakes or reservoirs.

Municipal wastewater treatment plants in many areas are required to remove phosphorous in their treatment process. While the biological treatment process removes some phosphorus, in most cases precipitation as an insoluble metal phosphate is required to meet discharge regulations. This precipitation step is normally accomplished with a metallic salt such as ferric sulfate, ferric chloride or aluminum sulfate. This precipitation step may be accomplished in the primary or secondary clarifiers.

Phosphorus Quantification Techniques Quantification of phosphorous requires the conversion of the phosphorus to

dissolved orthophosphate followed by colorimetric determination of dissolved orthophosphate. The analysis of different phosphorous forms (e.g. particulate or organic-P) is obtained by various pretreatment steps. Pretreatment may consist of filtering to remove suspended matter or various digestion techniques designed to oxidize organic matter.

Phosphorus can be present in surface waters as organic phosphorus, orthophosphate (an inorganic form of PO4), or as condensed (solid) phosphates. The phosphorus may be in solution or as a component of suspended particulates. The wet chemical colorimetric analysis of phosphorus only works for orthophosphates and thus other forms of phosphorus must be converted to this form if they are to be analyzed. Organic phosphorus can be oxidized (digested) using perchloric acid, nitric acid-sulfuric acid, or persulfate with the persulfate technique being the safest and least time consuming. The digestion methods are detailed in APHA method 4500-P B.

65


Three techniques for colorimetric analysis of phosphorus are available. The technique most commonly used is the ascorbic acid method, which can determine concentrations of orthophosphate in most waters and wastewater in the range from 2-200 µg P/L. Ammonium molybdate and antimony potassium tartrate react in an acid medium with dilute solutions of orthophosphate-phosphorus to form an intensely colored antimony-phospho-molybdate complex. This complex is reduced to an intensely blue-colored complex by ascorbic acid. The color is proportional to the phosphorus concentration. The complex is not stable and thus analysis must be performed within 30 minutes of adding the ammonium molybdate and antimony potassium tartrate.

Barium, lead, and silver interfere by forming a precipitate. The interference from silica, which forms a pale-blue complex is small and can usually be considered negligible. Arsenate is determined similarly to phosphorus and should be considered when present in concentrations higher than phosphorus.

Method Detection Limit "Method detection limit" is the smallest concentration that can be detected above

the noise in a procedure and within a stated confidence level. Several types of detection limits are used including instrument detection limit (IDL), method detection limit (MDL), and practical quantitation limit (PQL). The IDL is strictly instrument noise and does not include variability due to sample preparation steps. The MDL includes both instrument noise and sample preparation variability. The MDL is obtained by making a standard that is near the MDL and dividing it into at least 7 portions. Each of the portions is then processed through all sample preparation steps and then analyzed. The MDL is calculated using the following equation.

1,nMDL st α−= 6.1

where n is the sample size and α=0.01 is generally the required confidence. The student t distribution function is available in Excel as a two sided test statistic (so use TINV(2α,n-1)) and the standard deviation, s, can be computed in Excel as STDEV().

The PQL is about five times the MDL and represents a practical and routinely achievable detection limit with reasonable assurance that any reported value greater than the PQL is reliable. According to Standard Methods the method detection limit for phosphorus when using a 1 cm light path is approximately 150 µg P/L.

Spectrophotometer Limitations The diode array spectrophotometer has 316 diodes that cover the wavelength range

of 190 nm to 820 nm. Each diode generates a voltage output that is proportional to the number of incident photons. The voltage is then digitized, but the manufacturer of the instrument in the Cornell Environmental Laboratory, Hewlett-Packard, doesn't report the resolution of the analog to digital converter. At very low concentrations the difference between the intensity of light transmitted through the reference and the intensity of light transmitted through the sample approaches zero. At some low concentration the difference in light intensity approaches the resolution of the analog to digital converter. Another source of instrument error is drift in lamp intensity over

66


time. The lamp intensity is measured when a reference sample is made. The light intensity recorded by the diodes will vary proportionally to any lamp intensity drift.

The IDL should decrease as the number of diodes used in the analysis increases (as in Spectral analysis) for the same reason that replicate analysis of samples decreases the standard deviation. The "Spectral analysis" feature, which can be used to measure either single or multiple components, uses as much of the spectrum as the user desires and thus potentially decreases the IDL. Spectral analysis uses general least squares regression to add multiples of extinction coefficient arrays for each component to obtain the best curve fit for the sample. The extinction coefficient arrays are obtained from the slope of the linear regression line for absorbance as a function of concentration at each wavelength.

Experimental Objectives 1) Measure the concentration of phosphorus in several samples to test the precision

of the ascorbic acid technique. 2) Compare the results obtained using conventional analysis at a single wavelength

with spectral analysis. 3) Analyze the data using spectrophotometer software outside the lab. 4) Analyze multiple samples so that confidence intervals can be calculated. 5) Estimate the method detection limit (MDL). 6) Discuss methods to improve the method detection limit. 7) Discuss method automation.


Standards Preparation Method

1) Use 100 µg P/L stock. 2) Use a digital pipet and prepare 1 mL of each standard. 3) Use E-pure water to dilute the 100 µg P/L stock.

Reagent Addition for Samples and Standards 1) Pipette 1 mL sample into a disposable microcuvet using a 1 mL digital pipette.

2) Add 160 µL combined reagent and mix thoroughly by swirling. 3) After at least 10 minutes but no more than 30 minutes, measure absorbance of

each sample using a reagent blank as the reference solution.

Samples and Standards to Prepare 1) Reagent blank to be used as reference samples.

2) Prepare 6 standards containing 0 (reagent blank), 1, 3, 10, 30, 100 µg P/L.

3) Prepare 6 additional 10 µg P/L standards.

67


4) Prepare 5 samples such as Cayuga Lake water, tap water, Chem lawn runoff, local creeks…

Spectrophotometer Method 1) Use Sample Cuvettes. (Make sure to orient all cuvettes with the arrow on the left

because the cuvettes are not symmetrical and have different absorbance when turned 180°.)

2) Use the reagent blank as the reference sample for all samples.

3) Use units of µg P/L. 4) Label all samples with descriptions that your classmates will understand! 5) Fill in the general description with your initials and a description of the type of

samples

Samples Analysis

1) Measure the reference using a reagent blank. (The reagent blank is also the 0 mg/L standard.)

2) Analyze the reagent blank as a sample and verify that the absorbance deviates less than 0.004 AU (absorbance units) from zero. If the absorbance deviates more than 0.004 AU reanalyze the reference sample.

3) Analyze 6 standards as standards using the spectrophotometer and save the data as \\Enviro\enviro\Courses\453\phosphorus\netid_Pstd.

4) Analyze 6 standards as samples using the spectrophotometer and save the data as \\Enviro\enviro\Courses\453\phosphorus\netid_Pstdsam.

5) Analyze 7 10-µg P/L standards as samples and save the data as \\Enviro\enviro\Courses\453\phosphorus\netid_10Pstdsam.

6) Analyze 5 samples as samples using the spectrophotometer and save the data as \\Enviro\enviro\Courses\453\phosphorus\netid_Psam.

Prelab Questions 1) You will be creating 1 mL standards by diluting a stock of 100 µg P/L. Create a

table showing how you will prepare 1 mL of each of the standards. 2) All of the samples are diluted with a small amount of combined reagent. How is

this dilution accounted for when calculating the concentration of samples?

Data Analysis 1) Plot the absorbance spectra of the standards. What is happening in the UV region?

Are there any absorbance peaks? 2) Choose an appropriate wavelength (perhaps an absorbance peak) and use Excel to

create a calibration curve. For the calibration curve, absorbance should be a function of phosphorus concentration.

68


3) Use the 10-µg/L standards that were analyzed as samples to evaluate the Method Detection Limit using single wavelength analysis. Use your Spreadsheet to calculate the concentration of each of the replicates.

4) Use the 10-µg/L standards that were analyzed as samples to evaluate the Method Detection Limit using spectral analysis. Report the wavelength range used for your analysis. You will need to analyze each sample and copy the results into a spreadsheet. (Note that within a data file you change which sample the software is analyzing with the sample number control.)

5) Use the method with the lowest MDL to analyze your samples. Create a bar graph showing the concentration of each of the samples. Report the MDL in the figure caption.

Spreadsheet requirements Your spreadsheet must contain all of the analysis requested above as well as the following capabilities: 1) A well-marked cell containing the analytical wavelength for single wavelength

analysis. Changes to this cell must be reflected in all calculations and graphs. 2) The graph showing the absorbance spectra of the standards must have a vertical

line indicating the analytical wavelength. 3) All of the graphs must be on the same page as the analytical wavelength control

so the effect of changing the wavelength can be easily observed. 4) A separate sheet where you answer the 4 questions.

Hints If you haven't already learned how to use Vlookup() now is the time! The row() function returns the number of the row. I found it useful for this analysis! The slope() and intercept() functions eliminate the need to type equations off of graphs!

Questions 1) Which analysis technique gave the best results? Explain why. If the analytical

technique didn't significantly affect the MDL, explain why not. 2) What types of errors dominated your ability to measure phosphorus? 3) What method modifications do you propose to improve phosphorus

measurements? 4) Total phosphorus concentration in Cayuga Lake varies between 10 and 50 ppb.

Would the techniques used in this lab be able to measure these phosphorus levels?

References http://wwwrvares.er.usgs.gov/nawqa/circ-1136/h6.html#PHOS http://www.epa.gov/glnpo/lmmb/methods/index.html#Volume 3 Standard Methods for the Examination of Water and Wastewater.

69


Lab Prep Notes

Reagents

A Sulfuric acid solution, 4.9 N: Add 136 mL concentrated H2SO4 to 800 mL E-pure water. Cool and dilute to 1 L with E-pure water.

B Ammonium molybdate solution: Dissolve 40 g of (NH4 )6 Mo7O24•4H2O in 900 mL E-pure water and dilute to 1 L. Store at 4°C.

C Ascorbic acid: Dissolve 9 g of ascorbic acid (C6H8O6) in 400 mL E-pure water and dilute to 500 mL. Store at 4°C. Keep well stoppered. Prepare fresh monthly or as needed.

D Antimony potassium tartrate: Dissolve 3.0 g of K(SbO)C4H4O6•½H2O in 800 mL E-pure water and dilute to 1 L. Store at 4°C.

Combined color reagent: Combine the following solutions in order,

mixing (but do not entrain air as oxygen oxides the ascorbic acid) after each addition: (Prepare fresh weekly. Store at 4°C) Stock A, (4.9 N H2SO4) 50 mL Stock B, (Ammonium molybdate solution) 15 mL Stock C, (Ascorbic acid solution) 30 mL Stock D, (Antimony-tartrate solution) 5 mL

Water diluent solution: Add 4.0 g sodium lauryl sulfate and 5 g NaCl per L of E-pure water.

Stock phosphorus standard: Dissolve 0.4394 g of Potassium phosphate monobasic (KH2PO4) (dried at 105°C for one hour) in 900 mL E-pure water. Add 2 mL of concentrated H2SO4 and dilute to 1 L. 1.0 mL = 0.100 mg P (100 mg P/L).

Standard phosphorus solution: Dilute 1 mL of stock solution to 1 L. (1.0 mL = 0.1 µg P) (100 µg P/L).

Table 1. Reagents

Description Supplier Catalog

number concentrated

H2SO4 Fisher Scientific

(NH4 )6 Mo7O24•4H2O

Fisher Scientific

C6H8O6 Fisher Scientific K(SbO)C4H4O6•

½H2O Fisher Scientific

sodium lauryl sulfate

KH2PO4 Fisher Scientific


Description Supplier Catalog # 100-1095 µL

pipette Fisher Scientific 13-707-5

10-109.5 µL pipette Fisher Scientific 13-707-3 Disposable cuvets Fisher Scientific 14-385-942

Cuvet holder Fisher Scientific 14-385-939 UV-Vis

spectrophotometer Hewlett-Packard

Company 8452A

70


Soil Washing to Remove Mixed Wastes

Objective The goal of this laboratory exercise is to acquaint students with some of the

chemical reactions that result in the binding of inorganic and organic pollutants in subsurface materials. Extractants used by engineers to release contaminants at hazardous waste sites (where mixtures of both types of contaminants are present) may or may not prove effective, depending upon their mechanism of action. In this laboratory exercise, students will test the efficacy of a variety of proposed extractants in the removal of a mixture of an inorganic metal cation, and an organic compound from a contaminated porous medium.

Introduction Many Superfund site soils are contaminated with a mixture of contaminants

including toxic metals and organic compounds. A pressing environmental problem is to devise clean-up strategies that can effectively remove mixed wastes. Many kinds of contaminants bind to soils and aquifer media (collectively referred to here as porous media). Binding reactions limit the effectiveness of “pump and treat remediation” in which a contaminated porous medium is flushed with water to remove contaminants. In such cases, it can prove useful to engineer the properties of the aqueous phase to improve the mobility of the pollutants of interest.

In the case of toxic metals, release of medium-bound or “adsorbed” metals can be enhanced by introduction to the pore solution of a dissolved compound that will bind to the metal in the aqueous phase and form a dissolved “complex”. Such compounds are referred to as “ligands”, and ligands that bind metals very strongly are called “chelating agents”. Metal solubility and adsorption can also be strongly influenced by the oxidation state of the metal, and use of oxidants or reductants to alter the redox conditions in a porous medium can modify metal mobility both directly and indirectly. Direct effects would be observed if oxidized and reduced metal species have different adsorption characteristics (ex. Cr2O7

-2 vs. Cr+3). Indirect effects would be observed if a metal were bound to a solid phase that would be dissolved under different redox conditions (ex. Fe(OH)3 may dissolve under reducing conditions). Addition of acids or bases could also alter metal mobility. Adsorption of metals is very sensitive to pH shifts, with a decrease in pH favoring the release of cationic metal species (ex. Cd+2, Pb+2) and an increase in pH favoring release of anionic species (ex. Cr2O7

-2, SeO3-2).

Organic cations and anions will have a pH dependent adsorption behavior similar to that described above for metal ions. However, many organic pollutants of interest are nonionic and their binding to the matrix of the porous medium is not greatly influenced by pH. “Hydrophobic interactions” of nonionic organic compounds with organic matter in porous media appear to be a major driving force for their binding. The addition of surfactants to the pore solution can help to release sorbed nonionic organic pollutants. Under suitable conditions many organic pollutants can be degraded by addition of oxidants or by indigenous or added bacteria [i.e.; given that

71


the bacteria have the necessary genetic capabilities, nutrients (N, P, etc.) and a suitable electron acceptor]. Metals, however, are elements and cannot be degraded.

Porous media is not inert. The mineral and organic constituents of the porous matrix can react with added ligands, acids, bases, oxidants, reductants, and surfactants. A consequence, in some cases, is that a desired addition may be rendered impractical.

Given the variability and possible dissimilarity of conditions that influence the mobility of metal vs. organic pollutants, it is a challenging task to identify a remediation strategy that will successfully treat a given medium that is contaminated with mixed wastes. In this laboratory exercise, students will evaluate the utility of several alternative extractants for remediation of a soil that is contaminated with both a metal cation and an organic compound.

Theory

Binding Reactions The binding reactions of pollutants to the porous matrix may be classified, at least

in part, by where and how the binding reaction takes place. The term “adsorption” is used for reactions that take place at the interface between the solid and the solution. All other factors being equal, solids with a greater specific surface area (ex. units: m2/gram) will adsorb greater amounts of a dissolved solute. In adsorption reactions, the surface is referred to as an “adsorbent” and the solute as an “adsorbate”. Some adsorption reactions are driven by electrostatic attraction between the surface and the solute. “Ion exchange” is the term used for this type of reaction. All other factors being equal, surfaces with a greater number of charged sites per unit surface area will be able to bind greater quantities of dissolved ions. The concentration of surface exchange sites is commonly quantified as an “ion exchange capacity”. Surfaces with a high density of negatively charged sites (cation exchangers) will selectively bind positively charged ions while those with a high density of positively charged sites will be selective for anions.

“Absorption” is a process in which a solute penetrates within the solid matrix. “Partitioning” is a term that is synonymous with absorption. As an example, we would carry out a partitioning process if we were to add a pollutant to a separatory funnel containing water and an organic liquid such as octanol and then observe the resulting distribution of the contaminant between the aqueous and octanol liquid phases. The distributed contaminant would exist as a dissolved solute in each phase. As is noted below, the phase distribution behavior of nonionic organic pollutants in soils and aquifer media displays many characteristics of absorption reactions. The absorption of nonionic organics appears to be primarily into the organic matter content of the porous medium. This reaction is driven by the water loving nature of the solute, or lack thereof (i.e., pollutant “hydrophobicity”). All other factors being equal, porous media with higher organic carbon contents would have greater uptake of nonionic organic pollutants.

The term “sorption” is somewhat loosely used when the exact mechanistic nature of the pollutant’s distribution between the solution and the porous medium is not

72


understood, or when both adsorption and absorption reactions may contribute to the contaminant’s phase distribution.

Contaminant sorption reactions result from an reaction between a material that is dissolved in an aqueous solution with a solid phase. The physical/chemical properties of the contaminant, the solution and the sorbent all influence the resulting contaminant phase distribution. These influences are discussed below.

Sorbent Surface Charge As noted above, if the sorbate is an ion, then electrostatic attraction to the surface

can play an important role in contaminant adsorption. Virtually all soil surfaces are charged.

Oxide Minerals Surface charge can result from the ionization of surface functional groups in

response to the hydrogen ion concentration of the aqueous phase. Oxide minerals are often modeled as diprotic acids (Westall and Hohl, 1980). Accordingly the surface may donate two hydrogen ions as indicated by the following reactions:

1K2SOH SOH H+ +→ + 7.1

2KSOH SO H− +→ + 7.2

where SO represents the oxide surface that may exchange two hydrogen ions, and K1 and K2 are equilibrium constants for the first and second acid dissociation reactions.

Note, each dissociation constant can be thought of as a expression of the relationship between the concentration of protonated and deprotonated surface sites and the solution hydrogen ion concentration. Accordingly:

1

2

SOH HK

SOH

+

+

=

7.3

2

SO HK

SOH

− + =

7.4

K1 therefore represents the solution hydrogen ion concentration at which the

concentration of positively charged, diprotic, surface sites 2SOH+ is equal to that of

surface sites containing a single proton SOH . Similarly, when H+ equals K2 then

SO−

= SOH .

Although other models of the acid base behavior of oxide surface are conceivable, the above model is helpful in that it predicts that the surfaces can have both positively and negatively charged sites. With this model, H+ release from the surface will occur

73


in response to a decrease in the solution H+ concentration (i.e., an increase in pH, where pH is defined as -log H+ ). Accordingly, we would expect increasingly

higher solution pH conditions to favor formation of negatively charged surfaces, and this is observed. Different surfaces would have different acidity constants (K1 and K2) and would be expected to have different surface charges at the same solution pH. Each surface, at one unique pH, would have an equal concentration of 2SOH + and

SO− sites and would have no net charge. This is also observed and is referred to as

the pH point of zero charge (PZC). SiO 2, a common oxide in porous media (the main component of sand), has a low PZC (˜ pH 2 to 3) while iron and aluminum oxides (that commonly occur as surface coatings) have considerably higher PZCs (̃ pH 7 to 8) (Parks and DeBruyn, 1962).

Soil Organic Matter Another pH-dependent origin of surface charge is the ionization of the acidic

functional groups in soil organic matter. The carboxyl groups of humic-type organic matter typically have acidity constants = 10-5 (pK = 5) and are therefore highly ionized at circumneutral pH.

Isomorphic Substitution A final source of charge in soil is isomorphic substitution in the crystalline lattice

of some clay minerals. Substitution of Al+3 for Si+4 and Mg+2 for Al+3 will result in a net negative charge for the clay mineral phase.

The combined effects of isomorphic substitution, ionization of organic functional groups and the low PZC of silicon oxide minerals make it likely that many porous media will have a net negative charge. Consequently, stronger binding of cationic contaminants is generally anticipated.

Sorbent Ion Exchange Reactions Ion exchange reactions involve the exchange of ions of the same charge at an

oppositely charge site on the solid surface. Exchange reactions are often characterized by “selectivity coefficients” that may be thought of as equilibrium constants for the exchange reaction. For example, in the exchange of two monovalent cations, the exchange reaction may be depicted as:

yxKSO x y SO y x

− −+ + + +− + → − + 7.5

where:

yx

SO y xK

SO x y

− + +

− + +

− ⋅ = − ⋅

7.6

The magnitude of the selectivity coefficient, yxK , reflects the extent to which ion x+

vs. y+ will accumulated at the surface. Ions with high selectivity coefficients can displace more weakly held ions from an exchange site.

74


In a negatively charged soil, anionic compounds (ex. ionized organic acids, NO3-,

Cr2O7-2, etc.) will be repelled from the surface and therefore may be highly mobile.

Cationic species (ex. quaternary ammonium organic compounds, divalent transition metals, etc.) will be attracted to the surface and have restricted mobility. In principle, exchangeable pollutant cations may be mobilized by introduction of high concentrations of an innocuous cation. The practicality of such an approach would be dictated by the extent to which other exchangeable cations (that are not of environmental concern) are also exchanged. Since cations such as Na+, K+, Ca+2 Mg+2 are abundant in porous media, the amount of a cation added for exchange of a trace pollutant would have to be in great excess of the pollutant cation. As a result, release of contaminant cations by an ion exchange mechanism does not appear to be economically feasible.

Sorbent Hydrophobic Interactions The mechanisms responsible for the adsorption of charged species differ

considerably from those for nonionic compounds. Adsorption of charged ions may, in some cases, involve more than the simple electrostatic attraction of ions to a surface of opposite charge. Transition metal cations, for example, will often adsorb to oxide surfaces even under solution conditions that confer a positive charge on the surface (see additional discussion below under the topic of solution characteristics).

The sorption of nonionic organic pollutants behaves as if it is a partitioning process into the organic matter that is present as part of the soil matrix. Some of the general characteristics that lead to this conclusion are the observance of linear sorption isotherms at high solution concentrations (that can approach the solubility limit of solute compounds). [Note, an “isotherm” is simply the relationship between the quantity of pollutant that is bound (per unit mass or unit surface area of the sorbent) and the concentration of contaminant in solution.] In contrast, adsorption reactions are limited by the availability of surface sites and adsorption isotherms are typically non-linear at high solute concentrations. Partition reactions are also relatively free from competition (i.e., the presence of a second solute does not effect the sorptive uptake of the first) while competition for surface sites is an expected characteristic in an adsorption process. The extent of sorption of a given nonionic organic onto a variety of sorbents is highly correlated with their organic content as expressed by the weight fraction of organic carbon, foc (Karickhoff, 1984). For the same sorbent, the sorption of different nonionic solutes is highly correlated with their octanol-water partition coefficients (Kow) (Karickhoff, 1984). Collectively, these observations lead to the conclusion that the sorption of nonionic organic pollutants is primarily driven by hydrophobic interactions between the solute and the organic matter in the sorbent.

Solution pH Solution conditions can have dramatic effects on the adsorption of cationic

contaminants. For example the adsorption of cationic transition metals to oxide surfaces typically increases markedly over a narrow range of 1 to 2 pH units referred to as the “adsorption edge”. The pH dependence of metal ion adsorption can be explicitly accounted for by writing the adsorption reaction as:

75


xdKz z xx aqSOH Me SOMe H+ − ++ → + 7.7

where K d is the pH-dependent metal distribution coefficient, and according to Honeyman and Santschi (1988)

x

z x

d zx aq

SOMe HK

SOH Me

− +

+

⋅ = ⋅

7.8

A plot of logz x

zx aq

SOMe

SOH Me

−

+

⋅

versus pH, is referred to as a “Kurbatov plot”

(after Kurbatov et. al., 1951), and may be used to reveal the magnitude of the

exponent, x for H+ in the distribution coefficient. The ratioz x

x

SOMe

SOH

−

is the

quantity of adsorbed metal per unit surface. Since the above reaction and its equilibrium constant, Kd, are an over simplification of the actual adsorption mechanism, measured values of x are rarely integers. Nevertheless, x values ranging from 1 to 2 are common for adsorption of metal cations on oxide surfaces and demonstrate the strong dependence of the adsorption processes on pH. For example, if x = 2, an increase of 1 pH unit would result in a 100 fold increase in the amount of bound metal per unit surface (at the same solution concentration of metal ion). In general, adsorbed metal cations will be released as a consequence of a decrease in solution pH. Since the surfaces in the porous medium also have acid/base properties, and because many porous media contain acid-reactive components (such as carbonate minerals) a very large acid dose may be required to effectively alter the pH of the pore water. For this reason, acid extraction of adsorbed metals may not always be feasible.

Metal-Ligand Complexes Another influence of solution conditions on metal adsorption is through the

reactions of metals with ligands to form complexes. In some cases, metal-ligand complexes adsorb weakly or not at all (ex. Cl- complexes of Cd and Hg), in other cases metal-ligand complexes may adsorb with a binding strength greater than that of the free metal (ex. organic complexes of Cu) (Benjamin and Leckie, 1982). Judicious selection of a ligand for introduction into a porous medium may, therefore, be used to accomplish the release of adsorbed cations. Added ligands may, in some cases, undergo exchange reactions with the porous media or react to form complexes with cations that are not of environmental concern. For this reason the dose of a ligand needed to effectively release adsorbed metals will vary with the composition of the porous media and ligand addition may not prove feasible in some cases.

Oxidants and Reductants Changing solution composition by the introduction of oxidizing or reducing agents

may accomplish the release of adsorbed metals. Iron oxides are strong metal binding

76


agents and may be solubilized by reduction from ferric (Fe III) to ferrous (Fe II) iron. Many transition metals (e.g. Cd, Co, Cu, Ni, Pb, Zn) will remain as divalent cations during such a shift in redox status, and may therefore simply re-absorb to another surface. In some cases, alteration of the media redox conditions may directly influence metal mobility. For example reducing conditions would favor the presence of a cationic form of chrome (Cr+3) over the more mobile anionic form (Cr2O7

-2). Addition of oxidants may therefore help to mobilize chrome, however the organic matter in soils and ferrous minerals will also react with added oxidants.

In a manner similar to the role of iron oxides, the organic matter in porous media can be responsible for the binding of metal cations. The reaction of an added oxidant with humic-type organic matter may therefore accomplish solubilization of some metals (Lion et al., 1982). Strong oxidants will also act to break down organic contaminants.

Hydroxyl Radicals One application that has been used for remediation of organic contaminated soils is

the introduction of Fenton’s reagent. Fenton’s reagent is a mixture of hydrogen peroxide and ferrous iron (Fe+2). These chemicals react to produce hydroxyl radicals3 (OH•) according to the following reaction:

+2 +3 -2 2Fe + H O Fe +OH +OH→ i 7.9

The hydroxyl radicals produced by Fenton’s reagent are highly reactive and can effectively degrade recalcitrant aromatic compounds by ring substitution followed by ring cleavage (Sedlak and Andren, 1991).

Surfactants Additions of surfactants may aid in the release of sorbed nonionic organic

pollutants. In the case of sorption reactions that are driven by hydrophobic interactions, surfactant additions can have two beneficial effects: 1) a decrease in the aqueous activity coefficient for the dissolved nonionic organic compound and 2) formation of micelles in the aqueous phase.

The effect of the aqueous activity coefficient can be illustrated by examination of the sorption isotherm for the organic pollutant. If the isotherm is linear, then we may write:

SL L K CγΓ = 7.10

where Γ is the mass of solute sorbed per mass of solid, SLK is the sorptive distribution

coefficient, CL is the aqueous concentration of the sorbate, and γ is the activity coefficient of the dissolved sorbate.

Surfactants may act to decrease the activity coefficient, γ, for a nonionic molecule increasing the concentration in the aqueous phase in equilibrium with a given adsorbed amount, Γ. Surfactants increase the solubility of nonionic molecules

3 Radicals contain an odd number of electrons.

77


because the hydrophobic-nonionic molecules adsorb to the long hydrocarbon group while the ionic sulfonic group provides high solubility (Figure 1).

However, since surfactants are surface-active, they may also sorb to the porous medium, increase its organic content, and consequently increase the sorption of a nonionic organic contaminant. High concentrations of water-soluble cosolvents such as methanol and acetone can also act to decrease the activity coefficient, γ, and act to solubilize sorbed nonionic organic compounds (Schwarzenbach et al., 1993).

Surfactant molecules can aggregate into micelles in which their polar functional groups are oriented towards the aqueous solvent and their non-polar tails are oriented inward toward each other. The space within the micelles therefore provides a hydrophobic refuge for nonionic contaminants (Edwards et al., 1991). Surfactants will form micelles at aqueous concentrations greater than their “critical micelle concentration” (CMC). Since, as noted above, surfactants will sorb at the surface of the porous media, a high dose of surfactant may be required in order to maintain an aqueous concentration greater than the CMC.

Bacterial Polymers Many of the solution modifications discussed above involve the addition of

synthetic agents to contaminated soil to accomplish the release of sorbed contaminants. Natural constituents that occur in soils and aquifers may also enhance contaminant transport (McCarthy and Zachara, 1989). Bacterial polymers naturally occur in soil solution and have well-documented metal binding properties. The presence of bacterial polymers may therefore act as a natural process by which metal mobility is enhanced (Chen et al., 1995). The extracellular polymers produced by bacteria are hetero-polysaccharides and have high molecular weight. Interestingly, these large molecules have also been show to be effective at binding nonionic organic pollutants and at enhancing their transport in aquifer materials (Dohse and Lion, 1994). In principle, bacterial polymers with suitable binding properties could be produced in engineered reactor systems and be applied to contaminated waste sites to enhance the mobility of metal and nonionic organic contaminant mixtures. The efficacy of this type of remediation process has yet to be determined.

Apparatus Students will apply a range of extractant types (or mixtures of different types) to

remove contaminants (Zn and methylene blue) from a porous medium. Laboratory extractions will mimic an engineered soil washing system in which the contaminated soil is actively mixed with the extractant and then separated. A rotator will be used to provide agitation of samples of the medium with extractants, and a centrifuge will be used to provide phase separation. A UV/visible spectrophotometer with a diode array detector will be used to measure the concentration of the extracted organic pollutant.

R S

O

O

OO -

Figure 1. Molecular structure of detergent. The hydrocarbon chain (R group) of the detergent used in this lab is C12H25.

78


Extracted metal concentrations will be measured with an atomic absorption (AA) spectrophotometer.

Experimental Procedures Each group will develop their own hypothesis and experimental protocol. Different

concentrations of extractants, different organic contaminants, and different washing techniques could be the investigation subjects. Alternate organic contaminants should be cleared with the instructor prior to the lab period. Each group should limit the investigation to approximately 10 samples and should include appropriate controls and replicates.

The following protocol assumes that a common sand is employed to represent the porous medium. It is desirable, but not essential, to characterize each medium to be used (prior to the laboratory exercise) with respect to its carbon content [the “Walkley Black” method is one common procedure (Allison, 1965)], cation exchange capacity, and specific surface area [by sorption of ethylene glycol monoethyl ether (EGME) (Cihacek and Bremmer, 1979)].

I. Creation of a Contaminated Porous Medium A stock solution containing the soil contaminants will be provided [50 mg/L Zn

and 100 mg/L methylene blue]. For each extractant used in part II below, 2 samples of contaminated sand and one sample of clean sand will be used. The following procedure is based on the assumption that each student group will evaluate 3 extractants or 3 concentrations of an extractant. 1) Weigh out 9 aliquots of sand, 2.5±0.05 g each, and pour into 10 mL plastic

centrifuge tube. 2) Record the mass of the centrifuge tube with the sand (see Table 1). 3) Add 5 mL of the contaminant stock solution to 6 of the samples. 4) Add 5 mL distilled water to 3 of the samples (clean controls). 5) Place all of the samples on a rotator to mix the sand and the contaminant/clean

solutions. Agitate for 15 minutes. 6) Centrifuge the suspensions at 3000 x g for 5 minutes. 7) Pour the supernatant from the 6 contaminated sand samples into a 125 mL bottle. 8) Pour the supernatant from the 3 clean sand samples into a separate 125 mL bottle. 9) Weigh the centrifuge tubes with the sand and pore water. Calculate the volume of

pore water by subtracting the centrifuge tube and sand masses.

II. Determination of the Amount of Contaminant Sorbed by the Sand

Methylene blue - UV/Vis Spectroscopy Nitrate absorbs ultraviolet light and is present in the contaminated samples from

the addition of Zn(NO3)2·6H2O. We could account for this either by preparing a nitrate standard and using it as a component in spectral analysis or by eliminating the ultraviolet part of the spectrum from the analysis. We will eliminate the nitrate

79


interference by using a wavelength of 660 nm when measuring methylene blue. See page 160 for instructions on using the UV/Vis Spectrophotometer. 1) Measure the absorbance of 1, 5, and 10 mg/L methylene blue solutions as

“Standards.” Save the file as \\Enviro\enviro\Courses\453\soilwash\netid_MBstd.

2) Measure the absorbance of the combined supernatant from the 3 clean sand samples, the combined supernatant from the 6 contaminated sand samples, and the contaminating solution (diluted by a factor of 10) as “Samples.” Save the file as \\Enviro\enviro\Courses\453\soilwash\netid_contamsuper.

3) Record the concentration of methylene blue in the clean supernatant and contaminated supernatant (see Table 2). You can drag the blue cursor on the "standard graph" to the wavelength of choice and read the exact absorbance (and wavelength) in the digital display to the left of the graph and concentration in the digital display at the bottom of the Spectrophotometer window. If the clean supernatant has significant absorbance at 660 nm then alternate analytical techniques may need to be used.

4) The difference between the methylene blue concentration in the contaminant solution and the concentration in the supernatant may be used to determine the sorbed contaminant concentration as:

( )(solution volume)

=mass of sand

initial finalC CΓ 7.11

where Cinitial is the contaminant solution concentration and Cfinal is the concentration of the supernatant. Solution volume is the volume of contaminant added initially.

Zinc - Atomic Absorption Spectroscopy 1) Calibrate the AA using the zinc standards (1, 2, and 6 mg/L). 2) Dilute all of the following samples by a factor of 10 to ensure sufficient sample

volume for the analysis and to ensure that the results are in the calibrated range. 3) Measure and record the zinc concentration of the combined supernatant from the

3 clean sand samples. 4) Measure and record the zinc concentration of the combined supernatant from the

6 contaminated sand samples (see Table 4). 5) Measure and record the zinc concentration in the contaminating solution (the zinc

concentration should be close to 50 mg/L). 6) The difference between the zinc concentration in the contaminant stock and the

concentration in the supernatant may be used to determine the sorbed contaminant concentration using equation 7.11.

III. Soil Washing Solutions Students may wish to experiment with extractant mixtures. Some combinations of

extractant solutions may react violently! All proposed mixtures of extractants should

80


be cleared with the course instructor prior to their use. The following combinations should be avoided: mixtures of oxidants with reductants, mixtures of acids with bases, and mixtures of oxidants with organic extractants including: surfactants, chelating agents or cosolvents. The following extractant solutions will be available for testing. At least one group should measure the extractant capabilities of distilled water because it is by far the cheapest! 1) Distilled water. 2) Acid: ˜ 1 M solution of HCl prepared by diluting 27.4 mL of the concentrated

acid to 1 L with distilled water. 3) Cosolvent: 1:1 (v/v) mixture of acetone and distilled water. 4) Non-ionic surfactant: 10% (v/v) solution of Triton X-100 prepared by diluting

100 mL to 1 L with distilled water. Note: Triton X-100 is a non-ionic surfactant with a CMC of 2x10-5 M (Edwards et al., 1991). The chemical formula for Triton X-100 is:

(OCH 2CH 2)xOHCH3 -C-CH2 -C

CH3CH3

CH3CH3 where: x = 9 to 10, giving the surfactant a molecular weight of ˜ 607g/mole.

5) Anionic surfactant: 100 mM solution of dodecyl sulfate, sodium salt (C12H25SO4Na with MW of 288.4 so 28.84 g/L). This extractant works very well at full strength; lower concentrations could be investigated.

6) Chelating agent: ˜ 0.1 M solution of ethylenediamine-tetraacetate (EDTA) prepared by dissolving 37.22 g of the disodium salt in distilled water and diluting to 1 L.

7) Base: ˜ 1 M NaOH solution prepared by dissolving 40 g of NaOH distilled water and diluting to 1 L.

8) Oxidant: 1:1 (v/v) mixture of 30% H2O2 and distilled water.

9) Reductant: ˜ 1 M solution of Na2S2O3.5H2O prepared by dissolving 248 g in

distilled water and diluting to 1 L.

IV. Soil Washing Protocol The ability of each extractant to remove the zinc and methylene blue from the

contaminated soil will be measured by exposing the contaminated soil to the extractant and then measuring the concentration of the zinc and methylene blue in the extractant. 1) Add 5 mL of each extractant to be tested to 2 contaminated sand samples and 1

clean sand sample. 2) Place the sand extractant mixtures on a rotator to mix for 15 minutes. 3) Centrifuge the suspensions at 3000 x g for 5 minutes. 4) Decant the supernatant from each centrifuge tube into labeled 15 mL bottles. A

small air line can be used to help force the supernatant from the centrifuge tubes.

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V. Analysis of Extracted Metal and Organic Pollutants

Methylene blue - UV/Vis Spectroscopy Note that it is unnecessary to measure the methylene blue concentration in samples

that do not appear to have any blue. Samples that are visually free of methylene blue can be recorded as 0 mg/L methylene blue. 1) Measure the absorbance of each of the clean sand extracts as “Samples.” All of

the clean extracts can be analyzed together if desired. Save as \\Enviro\enviro\Courses\453\soilwash\netid_cleanext.

2) Measure concentration of methylene blue in each of the clean extracts based on the absorbance at 660 nm (see Table 2).

3) Measure the absorbance of each of the extracts of the contaminated sand as “unknowns.” There will be 2 replicates for each extract. All of the contaminated extracts can be analyzed as a group so that their results are saved in a single file. Save as \\Enviro\enviro\Courses\453\soilwash\netid_contamext.

4) Measure concentration of methylene blue in each of the contaminated extracts based on the absorbance at 660 nm (see Table 2). Note that it may be necessary to choose a different analytical wavelength or to dilute the sample if the absorbance exceeds ˜2.5 at 660 nm.

5) Calculate the mass extracted per mass of sand as:

( ) ( )extractedC solution volume

mass of sand∆Γ = 7.12

where solution volume is the sum of residual pore water volume after decanting the contaminant plus the extractant volume.

Zinc - Atomic Absorption Spectroscopy 1) Dilute all samples by a factor of 10 prior to analysis. 2) Measure and record the absorbance of the supernatant from the 3 clean sand

samples (see Table 4). 3) Measure and record the absorbance of the supernatant from the 6 contaminated

sand samples. Dilute the supernatant with distilled water if the absorbance is not less than the absorbance of the 6-mg/L standard.

4) Calculate the concentration of zinc in each of the sand extracts. 5) Calculate the mass of Zn removed per mass of sand using equation 7.12.

The results for any extractant must justify the cost of its use. The results obtained by extraction of contaminated soil using distilled water serve as a basis for comparison to which results obtained with extractants should be compared. Results from all extractants or extractant combinations evaluated should be compared to provide an inter-comparison of their relative effects of the removal of metal and organic pollutants.

82


Prelab Questions 1) The point of zero charge for SiO 2 is approximately at pH = 2.5. Is the charge of

SiO2 positive or negative at a pH of 7?

2) Do cations or anions generally bind most strongly to soil? 3) Develop a hypothesis concerning soil washing, and write an experiment protocol

to test your hypothesis that you can do in a lab period. Include detail of concentrations of extractants and contaminants for each vial. You may want to work with your lab partner(s) because this will be your experiment! Design your experiment to use no more than 9 vials.

Data Analysis 1) Report the contaminated sand concentration (grams of contaminant/gram of sand)

for zinc and methylene blue. 2) Calculate the fractional removal of zinc and methylene blue for each extractant or

extractant concentration. The fractional removal based on the amount of

contaminant initially sorbed is f = ∆ΓΓ

where Γ is defined in equation 7.11 and

?Γ is defined in equation 7.12. Present this using an appropriate graph. 3) Discuss which extractant performed best at removal of the zinc. Which was best

at removing the test organic? Which extractant worked best at removing both contaminants? Discuss these results in terms of the chemical change that the extractant was designed to accomplish. (Note that these questions may need to be modified based on the samples you analyzed.)

4) Discuss any difficulties in evaluating extractant effectiveness and propose improved analytical techniques.

5) Discuss your results and their implications for the hypothesis that you developed. 6) Analyze your results including the reproducibility of replicate analyses in terms of

possible sources of error. 7) Suggest options for additional research.

References Allison, L. E., “Organic carbon”, in: Soil Analysis Part 2: Chemical and

Microbiological Properties, C. A. Black (ed.), Amer. Soc. Agronomy, Madison, WI, p 1367, 1965.

Benjamin, M.M. and J.O. Leckie, “Effects of complexation by Cl, SO4 and S2O3 on adsorption behavior of Cd on oxide surfaces”, Env. Sci. & Tech. 16(3), pp. 162-170, 1982.

Chen, J.-H., L.W. Lion, W.C. Ghiorse, and M.L. Shuler, “Trace metal mobilization in soil by bacterial extracellular polymers”; Water Research, (1995, in press).

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Cihacek, L.J. and J.M. Bremmer, “A simplified ethylene glycol monoethyl ether procedure for assessment of soil surface area”, J. Soil Sci. Soc. Am. 43 pp. 821-822, 1979.

Dohse, D.M., L.W. Lion, “The effect of microbial polymers on the partitioning and transport of phenanthrene in a low-carbon sand”; Environmental Sci. & Tech. 28(4), 541-547 (1994).

Edwards, D.A., R.G. Luthy, and Z. Liu, “Solubilization of polycyclic aromatic hydrocarbons in micellar nonionic surfactant solutions”, Env. Sci. & Tech. 25, pp. 127-133, 1991.

Honeyman, B.D. and P.H. Santschi, “Metals in aquatic systems”, Env. Sci. & Tech. 22(8), pp. 862-871.

Karickhoff, S.W., "Organic Pollutant Sorption in Aquatic Systems", J. Hydraulic Engrg., 110(6), p. 707, 1984.

Kurbatov, M.H., G.B. Wood, and J.D. Kurbatov, “Isothermal adsorption of cobalt from dilute solutions”, J. Phys. Chem. 55, pp. 1170-1182, 1951.

Lion, L.W., R.S. Altmann, and J.O. Leckie, "Trace metal adsorption characteristics of estuarine particulate matter: Evaluation of contributions of Fe/Mn oxide and organic surface coatings," Env. Sci. and Tech. 16(10), pp. 660-666 (1982).

McCarthy, J. F. and J. M. Zachara, “Subsurface transport of contaminants”, Env. Sci. & Tech. 23, pp. 496-502, 1989.

Parks, G.A. and P.L. DeBruyn, “The zero point of charge of oxides”, J. Phys. Chem. 66, pp. 967-973, 1962.

Sedlak, D.L., and A.W. Andren, “Oxidation of chlorobenzene with Fenton’s reagent”, Env. Sci Tech. 25(4), pp. 777-782, 1991.

Schwarzenbach, R.P., P.M. Gschwend, and D.M. Imboden, Environmental Organic Chemistry, Wiley Interscience Publ., New York, NY; 681 pp., 1993.

Standard Methods for the Examination of Water and Wastewater, L.S. Clesceri, A.E. Greenberg and R.R. Trussell (eds.) 17th edition, Am. Public Health Assoc.(Publisher), Washington, DC; 1989.

Westall, J. and H. Hohl, “A comparison of electrostatic models for the oxide/solution interface”, Adv. Colloid Interface Sci. 12 p. 265, 1980.

84


Table 1. Data table.

bottle # contaminated or clean

mass sand (g)

mass bottle + sand (g)

mass bottle + sand + pore

water (g) clean cont. cont. clean cont. cont. clean cont. cont.

Table 2. Methylene blue data table.

concentration

clean sand supernatant contaminated sand supernatant

extractant 1 clean sand extractant 1 cont. sand rep 1

extractant 1 cont. sand rep 2

extractant 2 clean sand extractant 2 cont. sand rep 1 extractant 2 cont. sand rep 2


Table 3. Zinc concentration measurements data table.

dilution concentration

clean sand supernatant contaminated sand supernatant

extractant 1 clean sand

extractant 1 cont. sand rep 1

extractant 1 cont. sand rep 2 extractant 2 clean sand

extractant 2 cont. sand rep 1 extractant 2 cont. sand rep 2


85


Lab Prep Notes



Zn(NO3)2·6H2O Fisher Scientific Methylene Blue Fisher Scientific M291-25

HCl Fisher Scientific NaOH Fisher Scientific S318-500 H2O2 Fisher Scientific H325-500

Na2S2O3 Fisher Scientific S445-500 nitrilotriacetic

acid Aldrich N840-7

Triton X-100 Aldrich 23,472-9 acetone Fisher Scientific A929-1

FeSO4·7H2O Aldrich 31,007-7 alginic acid, sodium salt

Aldrich 18,094-7

Fe(NO3)3·9H2O Aldrich 21,682-8 Humic acid Aldrich H1,675-2

Nitric Acid 6 N Fisher Scientific LC17-70-2 Dodecyl sulfate,

sodium salt (C12H25SO4Na)

Aldrich 85,192-2

Zinc reference solution

Fisher Scientific SZ13-100

Table 5. Stock Solutions (100 mL each). Description MW (g/M) conc. (g/L) 100 mL

C16H18N3SCl 319.87 10 1 g Zn(NO3)2·6H2O 297.4 227.4048 22.74 g

5 g as Zn

Table 6. Contaminating Solution (1 L)

Description MW (g/M)

conc. (mg/L)

1 L

C16H18N3SCl 319.87 100 10 mL stock Zn(NO3)2·6H2O 297.48 50 (as Zn) 10 mL stock

86


Table 7. Equipment list Description Supplier Catalog

number refrigerated

centrifuge MP4R Fisher Scientific 05-006-4

4-place rotor 4B Fisher Scientific 05-006-9 Diode array

spectrophotometer Hewlett-Packard 8452A

rototorque rotator Cole Parmer H-07637-00 10 mL centrifuge

vials Fisher Scientific 05-529-1A

repipet II Dispensor Fisher Scientific 13-687-62B PP bottles 15 mL Fisher Scientific 02-923-8G

Zinc Disposal Guidelines The amount of zinc that can be disposed to the sanitary sewer is limited. The

wastewater treatment plant has a limit on the concentration of zinc that can be in the sludge. The Zinc stock solution should not be disposed to the sanitary sewer. Zinc that is sorbed to the sand can be dried and sent to the landfill in the trash.

Setup

1) Use repipet dispensors for contaminating solution, distilled water and possibly for additives.

2) Prepare calibration standards for the AA and for the UV-Vis spectrophotometers. 3) Each group needs 9 centrifuge vials, 20 15-mL bottles, and 2 125-mL bottles. 4) Connect a very fine tube to an air line at each island to be used to help empty the

supernatant from the centrifuge vials. 5) Devise technique to filter samples prior to AA analysis!

Class Plan

1) Demonstrate use of AA when samples contain particulate matter. (Keep sipper tube off of the bottom of the vial!)

2) EDTA works well for Zn at 0.1 M 3) Dodecyl sulfate works well for MB at 0.01 M

87

Oxygen Demand Concepts and Dissolved Oxygen Sag in Streams


Introduction In recent years “biodegradable” has become a popular word. Often it is assumed

that if something is biodegradable, then disposal is not a problem. We know that throwing non-biodegradable substances into our environment leads to degradation of our planet. But disposal of biodegradable compounds also can be detrimental to the environment.

The effects of improper disposal of biodegradable substances became a source of public outrage in the early 1800's. The flush toilet was becoming popular and sewage was discharged directly into the nearest waterway. The receiving waters were quickly polluted. Fish in the receiving waters died and the water had a very offensive odor. Although there are many reasons why we no longer discharge untreated sewage into the environment, (including disease transmission, sediment buildup...) one of the reasons is directly related to the fact that sewage contains much that is biodegradable.

Theory Biodegradable means that a substance can be converted into simpler compounds by

biologically mediated reactions. The second law of thermodynamics predicts that oxidation of high energy level organics (relative to low energy level CO2) is favored. Oxygen is one of the strongest oxidizing agents found in natural aquatic systems. Oxidation reactions are thermodynamically favored, but kinetically slow unless microbially mediated. The end products of complete aerobic biodegradation are CO2 and H2O. Production of CO2 has recently come under fire as a potential cause of global warming, but that is not the subject of this lab. The problem is not with the products of biodegradation, the problem is that aerobic biodegradation of a compound requires another reactant. Let's look at the biodegradation of a simple organic compound, glucose.

6 12 6 2 2C H O +? 6CO +6H O→ 8.1

To balance the equation oxygen is needed.

6 12 6 2 2 2C H O +6O 6CO +6H O→ 8.2

The consumption of oxygen needed for biodegradation can be a problem. Oxygen is not very soluble in water. The equilibrium concentration of oxygen in water is approximately 10 mg/L (see Figure 1). That means that the degradation of a few mg/L of a biodegradable compound in a river could result in the depletion of dissolved oxygen. Fish have a bad day when oxygen is depleted from their environment. Some species of fish such as trout begin to suffer when the dissolved oxygen concentration drops below 5 mg/L.

88


Oxygen in water is consumed during aerobic biodegradation of organic compounds and is replenished from the atmosphere. The two processes have different kinetics, but are coupled. As the oxygen is depleted by biodegradation the rate at which oxygen is transferred into the water increases because the transfer driving force increases. The rate at which oxygen is dissolved into water from the atmosphere is proportional to the deficit of oxygen in the water. The oxygen deficit is simply the difference between the equilibrium oxygen concentration and the actual oxygen concentration. These two reactions (reaeration, and biochemical utilization) are modeled by the Streeter-Phelps equation. In order to increase the rate at which the biodegradation occurs, the concentration of bacteria was increased for use in this laboratory experiment. Bacteria respire and thus consume oxygen even when no substrate is present. Thus an additional term for bacterial respiration will be needed to model the oxygen sag results obtained in the laboratory.

Streeter Phelps Equation Development We’ll begin by developing the oxygen deficit as a function of time in a completely

mixed batch reactor (no inflow and no outflow) with initial concentrations of Biochemical Oxygen Demand (BODL) and dissolved oxygen. We will include oxidation of BODL and reaeration from the atmosphere. These effects are coupled in equation 8.3 where C represents oxygen concentration. The first two terms on the right are negative since oxidation of BOD and respiration consume oxygen while the third term is usually positive since reaeration increases the concentration of oxygen (except in the rare instance where the dissolved oxygen concentration is greater than the equilibrium dissolved oxygen concentration). Eventually we will make a comparison between time in a reactor and distance down a river.

respirationoxidation reaerationCC CCt t t t

∂∂ ∂∂= + +

∂ ∂ ∂ ∂ 8.3

Oxidation of BOD We must first develop a relationship for the change in oxygen concentration due to

oxidation of organics. The rate that oxygen is used will be proportional to the rate that substrate (or biochemical oxygen demand) is oxidized. The rate of substrate utilization by bacteria is given by the Monod relationship

6789

101112

10 20 30 40

Temperature (°C)

Figure 1. Equilibrium dissolved oxygen concentration as a function of water temperature.

89


s

dL kLXdt K L

−=

+ 8.4

where L is substrate concentration expressed as oxygen demand or BODL [mg/L], k is the maximum specific substrate utilization rate, Ks is the half velocity constant, and X is the concentration of bacteria. However, the concentration of bacteria is a function of the substrate concentration and thus application of the Monod equation to a polluted river is not trivial. Often the bacterial concentration remains relatively constant. If the half velocity concentration is large relative to the concentration of substrate we obtain

oxs s

dL kXL kXL k L

dt K L K − −

= ≅ ≅ − + 8.5

where kox is a first order oxidation rate constant that includes both the approximation that the bacteria concentration is roughly constant and that the substrate concentration is smaller than the half velocity constant.

Separate variables and integrate

0

= ( )o

L t

oxL

dLk dt

L−∫ ∫ 8.6

to obtain

oxk toL L e−= 8.7

The rate of oxygen utilization is equal to the rate of substrate utilization (when measured as oxygen demand) and thus we have

ox= = -k LoxidationC dLt dt

∂∂

8.8

where C is the dissolved oxygen concentration [mg/L]. Now we can substitute for L in equation 8.8 using equation 8.7 to obtain

ox o= -k L e oxk toxidationCt

−∂∂

8.9

Respiration Bacteria utilize oxygen for respiration and for cell synthesis. When no substrate is

present the bacteria cease synthesis, but must continue respiration. This continual use of oxygen is termed "endogenous respiration." Bacteria use stored reserves for endogenous respiration. We can model this oxygen demand as a constant that is added to the demand for oxygen caused by substrate utilization. As a first approximation, we can assume that this oxygen demand is proportional to the concentration of bacteria. In addition, we will assume that the population of bacteria is relatively constant throughout the experiment.

90


e=-bX =-krespirationC

t

∂

∂ 8.10

where b is the specific endogenous oxygen consumption rate and ke is the endogenous oxygen consumption rate.

Oxygen Transfer Coefficient The rate of oxygen transfer is directly proportional to the difference between the

actual dissolved oxygen concentration and the equilibrium dissolved oxygen concentration.

( )*,

ˆ= C - Creaerationv l

Ck

t∂

∂ 8.11

where C* is the equilibrium oxygen concentration, C is the actual dissolved oxygen concentration, and ,

ˆv lk is the is the overall volumetric oxygen transfer coefficient. If

reaeration is the only process affecting the oxygen concentration then equation 8.11 can be integrated to obtain

*

, 0*0

ˆln ( )v lC C

k t tC C

−= −

− 8.12

Oxygen Deficit We now have equations for the reaction of oxygen with BODL, endogenous

respiration, and for reaeration. Substituting into equation 8.3 we get

( )*ox o e ,

ˆ= -k L k + C - Ce oxk tv l

Ck

t−∂

∂ 8.13

We can simplify the equation by defining oxygen deficit (D) as:

*D=C -C 8.14 and noting that the rate of change of the deficit must be equal and opposite to the rate of change of oxygen concentration

= dC dDdt dt

− 8.15

We must remember that the deficit can never be greater than the equilibrium concentration (D must always be less than C*)! In addition, the BOD model breaks down if the dissolved oxygen concentration is less than about 2 mg/L because the

lack of oxygen will limit microbial kinetics and dLdt

will no longer equal - oxk L . If we

stick to conditions under which our assumptions are valid then we can substitute equations 8.14 and 8.15 into equation 8.3 to obtain

,ˆ - o xk t

e ox o v l

Dk k L k D

te−∂

= +∂

8.16

91


This is a first order linear differential equation. Integration with initial oxygen deficit = Do @ t = 0 gives:

, ,ˆ ˆ-

, , ,

- ˆ ˆ ˆ

v l v loxk t k tk te e ox oo

v l v l v l ox

k k k LD D

k k k ke e e− −

= + + −

8.17

Application to a River We are interested in the oxygen deficit as a function of distance down a stream. As

an approximation we can think of a cross section of a river as a completely stirred reactor that is slowly moving downstream. The relation between time in a batch reactor and distance down the river is simply

t = xu

8.18

where u is the stream velocity and x is distance. The Streeter-Phelps model assumes a constant input of biodegradable substrate, Lo, at x = 0 and the model is valid under steady-state conditions.

Of particular concern is the maximum deficit, Dc. We want to know the value of Dc

and where (or when) it will occur ( ct = cxu

). This will be the "critical point." If there

are going to be adverse effects (like dead fish) this will be the place. The maximum oxygen deficit occurs when

= 0Dt

∂∂

8.19

We can substitute this into the first order differential equation 8.6 to get

,ˆ0 - o x ck t

e ox o v l ck k L k De−= + 8.20

and solve for Dc to get

,

ˆ

o x ck te ox o

c

v l

k k LD

k

e−+= 8.21

an equation with unknowns tc and Dc. The Streeter-Phelps equation still holds at the critical point so we also have

, ,ˆ ˆ-

, , ,

= - ˆ ˆ ˆ

v l c v l co x ck t k tk te e ox oc o

v l v l v l ox

k k k LD D

k k k ke e e− −

+ − + −

8.22

also with unknowns xc and Dc. So now we have two equations in two unknowns. We can solve for tc by eliminating Dc.

92


( ) ( ), , ,

2,

ˆ ˆ ˆ- -1= lnˆ -

e v l o v l ox v lc

oxo oxv l ox

k k D k k kt

kL kk k

+

8.23

To find Dc given the kinetic coefficients and the initial oxygen deficit, first find tc using equation 8.23. Then use equation 8.21 to solve for Dc.

Zero Order Kinetics An alternate model can be derived based on the assumptions that the concentration

of bacteria is relatively constant and that the rate of substrate utilization is zero order, i.e., the concentration of the substrate is greater than the half velocity constant Ks.

0= - =-s

dL kLXkX k

dt K L−

≅+

8.24

The change of the deficit of dissolved oxygen is equal to the change caused by microbial degradation (k0) plus change due to endogenous respiration minus the

reaeration ( ,ˆ

v lk D ).

0 ,ˆ= + e v l

dDk k k D

dt− 8.25

This equation is only valid when the substrate concentration is greater than Ks. To simplify derivation, assume that Ks is very small relative to the initial BOD added to the system and apply the zero-order model until the substrate is completely oxidized. When the substrate concentration reaches zero a discontinuity will occur as substrate oxidation stops. Separating variables and integrating

00 ,

= ˆ

o

D t

D e v l

dDdt

k k k D−+∫ ∫ 8.26

0 ,

, 0 ,

ˆ1ln =

ˆ ˆe v l

v l e v l o

k k k Dt

k k k k D

−

−

+−

+ 8.27

and solving for the dissolved oxygen deficit

,

ˆ

0 0 ,

,

ˆ( )=

ˆ

v ltke e v l o

v l

k k k k k DD

k

e−−+ +

8.28

The substrate concentration is depleted when 0

= oLt

k. Substituting into equation 8.28

to get the maximum dissolved oxygen deficit yields

93


,

0

ˆ

0 0 ,

,

ˆ( )=

ˆ

o v lL k

ke e v l o

t

v l

k k k k k DD

k

e −

−+ + 8.29

where Dt is the dissolved oxygen deficit at the transition when the substrate is all

utilized. For times greater than = oLt

k there is no longer any substrate and thus k0 = 0

and equation 8.25 becomes

,ˆ = e v l

dDk k D

dt− 8.30

Separating variables and integrating

,

= ˆ

t t

D t

D te v l

dDdt

k k D−∫ ∫ 8.31

? ,

, ,

ˆ-1ln = -

ˆ ˆe v l

t

v l e v l t

k k Dt t

k k k D

−

−

8.32

( ) ( ),

ˆ

,

,

ˆD=

ˆ

v l tk t te e v l t

v l

k k k D

k

e −−

8.33

Equation 8.33 is valid for all times greater than oLk

. The general shapes of the two

types of sag curves are shown in Figure 2.

Experimental Objectives The objectives of this lab are to:

1) Illustrate the effects of adding biodegradable compounds to natural waters.

2) Evaluate the Streeter-Phelps dissolved oxygen sag model and a zero order substrate utilization model and compare with laboratory data.

3) Explain the theory and use of dissolved oxygen probes.

6.0

0.0

1.0

2.0

3.0

4.0

5.0

800.00.0 200.0 400.0 600.0

Time (s)

DO sag

(mg/L)

zero order

first order

Figure 2. Dissolved oxygen sag curves obtained from zero and first order models for substrate utilization.

94


Experimental Methods In this lab we will examine the

effects of adding a small amount of a biodegradable compound to a small batch reactor. We will measure the dissolved oxygen concentration over time using a dissolved oxygen probe. The apparatus is shown in Figure 3.

The BOD measured using this technique will be lower than the BOD measured using the standard BOD test because a significant fraction of the glucose will be converted into cell material (i.e. used for synthesis instead of for respiration). This technique can be used to obtain kinetic parameters for yield, half velocity constant and maximum substrate utilization rate (Ellis et al., 1996).

Probe Calibration Calibrate the dissolved oxygen probe (see

http://www.cee.cornell.edu/mws/Software/DOcal.htm).

Oxygen Transfer Coefficient

1) Prepare to monitor dissolved oxygen. 2) Place the dissolved oxygen probe in the reactor. 3) Pour 50 mL of deoxygenated distilled water into the batch reactor. 4) Set the stirrer speed to 5. 5) Set the airflow rate to 50 mL/min. 6) Monitor the dissolved oxygen for 3 minutes (or longer). 7) Save the data as \\Enviro\enviro\Courses\453\oxygen\netid_O2trans. The data will

be used later to estimate the oxygen transfer coefficient.

Endogenous respiration oxygen requirements

1) Pour 50 mL of a bacterial suspension into the batch reactor. 2) Place the dissolved oxygen probe in the reactor. 3) Set the stirrer speed to 5. 4) Set the airflow rate to 250 mL/min to aerate the reactor contents. 5) Prepare to monitor dissolved oxygen.

DO probe

Hypodermic diffuser

100 mL beaker Water surface

Stirbar

Figure 3. Apparatus used to measure dissolved oxygen consumption rates.

95


6) After the dissolved oxygen concentration is close to saturation turn off the air and monitor the dissolved oxygen for 3 minutes (or longer).

7) Save the data as \\Enviro\enviro\Courses\453\oxygen\netid_endog. The data will be used later to estimate the endogenous respiration rate.

BOD of glucose solution

1) Set the airflow rate to 250 mL/min and aerate the bacterial suspension used previously.

2) Prepare pipette to add 75 µL of glucose solution (this will provide a BOD of 15 mg/L when diluted by 50 mL bacterial suspension).

3) Prepare to monitor dissolved oxygen. 4) Turn off the airflow. 5) As quickly as possible, add glucose through the port in the bottle and begin

monitoring the dissolved oxygen concentration. 6) Monitor the dissolved oxygen until the dissolved oxygen concentration reaches

approximately 0 mg/L. 7) Save the data as \\Enviro\enviro\Courses\453\oxygen\netid_BOD. The data will

be used later to estimate the BOD of the glucose solution.

DO Sag Curves

1) Set the airflow rate to 250 mL/min and aerate the bacterial suspension used previously.

2) Prepare to monitor dissolved oxygen. 3) Reduce the airflow rate to 50 mL/min. 4) Begin monitoring the dissolved oxygen in the reactor. Use 5 second data intervals.

5) After ˜300 seconds of monitoring add 10 mg glucose BOD/L (50 µL stock) to the reactor.

6) Observe the oxygen depletion in the reactor. 7) Continue monitoring until the dissolved oxygen concentration returns to within

90% of the original DO concentration. 8) Save the data as \\Enviro\enviro\Courses\453\oxygen\netid_sag.

Prelab Questions 1) A dissolved oxygen probe was placed in a small vial in such a way that the vial

was sealed. The water in the vial was sterile. Over a period of several hours the dissolved oxygen concentration gradually decreased to zero. Why? (You need to know how dissolved oxygen probes work to answer this!)

2) Which assumption is different between the Streeter-Phelps and the zero order model?

96


Data Analysis The rate constants can be estimated using Excel. A sample spreadsheet is available

at the course web site.

Oxygen Transfer Coefficient

1) Estimate the gas transfer coefficient from equation 8.12 or by using the spreadsheet model.

2) Graph the dissolved oxygen concentration vs. time along with the theoretical curve.

Endogenous Decay

1) Estimate the endogenous oxygen consumption rate from the slope of the graph or by using the spreadsheet model.

2) Graph the dissolved oxygen concentration for the bacteria culture in the BOD bottle without any added BOD vs. time along with the theoretical curve.

BOD of Glucose

1) Use the "DO sag" Excel spreadsheet to estimate the first or zero order oxygen utilization coefficients, and the BOD exerted by the glucose. Which model fits the data best?

2) How long did it take for the biodegradation of the glucose to occur? 3) What was the change in dissolved oxygen concentration during that time? 4) How much BOD did the glucose solution exert expressed as a fraction of the

BOD of the glucose added. 5) Graph the dissolved oxygen concentration vs. time for the glucose solutions along

with the theoretical curves. Identify the regions where biodegradation of the glucose was occurring.

Dissolved Oxygen Sag

1) Use the previous estimates of the oxygen transfer coefficient, endogenous respiration rate, and fraction of BOD exerted (note that a different amount of BOD was added for the sag curve than for the BOD measurement!) to plot zero and first order model predictions of the dissolved oxygen sag curve. Discuss any discrepancies.

2) Estimate the first and zero order oxygen utilization coefficients and the BOD exerted using the spreadsheet models by minimizing the RMSE using both models with your data. Use the endogenous respiration rate and the reaeration rate estimated previously. Which model (zero or first order) fits the data best? Are the fit parameters significantly different than those obtained in the BOD of glucose analysis? Include the estimated parameters in your report.

3) Graph the dissolved oxygen concentration vs. time for the dissolved oxygen sag curve along with the theoretical curves.

97


4) On the graph indicate maximum dissolved oxygen sag and compare with the BOD added.

5) Why is the dissolved oxygen sag less than the BOD added?

References Ellis, T. G.; D. S. Barbeau; B. F. Smets and C. P. L. J. Grady. 1996. “Respirometric

technique for determination of extant kinetic parameters describing biodegradation” Water Environment Research 68(5): 917-926.

98


Lab Prep Notes

Bacterial stock preparation using 20% PTYG

Grow 4 liter culture of Ps. putida 1) Heat 1 L of distilled water and

dissolve media for 4 L of 20% PTYG.

2) Dilute to 4 L in 6 L container containing aeration stone and stirrer.

3) Thaw one cryovial containing Ps. putida and transfer into PTYG media.

4) Stir and aerate for 24 hours.

Wash/enumerate Ps. putida culture 1) Centrifuge 4 L culture in 250 mL

bottles to obtain concentrated stock (5000 rpm for 10 minutes).

2) Resuspend total culture in 500 mL using 10x BOD dilution water (pH control is essential for bacterial growth and trace nutrients are required).

3) Refrigerate at 4°C.

Table 3. 20% PTYG culture media. (Prepare 4 L)

compound mg/L g/4L peptone 1000 4 tryptone 1000 4

yeast extract 2000 8 glucose 1000 4 MgSO4 470 1.9

CaCl2·2H2O 70 0.28

Setup

Table 1. Reagent list


peptone Fisher Scientific BP1420-100 tryptone Fisher Scientific BP1421-100 glucose Aldrich 15,896-8

yeast extract Fisher Scientific BP1422-100 MgSO4·7H2O Fisher Scientific CaCl2·2H2O Fisher Scientific

KH2PO4 Fisher Scientific K2HPO4 Fisher Scientific

Na2HPO4 · 7H2O

Fisher Scientific

NH4Cl Fisher Scientific FeCl3 · 6H2O Fisher Scientific



magnetic stirrer Fisher Scientific 11-500-7S Accumet™ 50


ATI Orion DO probe


6 L container Fisher Scientific 03-484-22 250 mL PP

bottle Fisher Scientific 02-925D

15 mL PP bottles

Fisher Scientific 02-923-8G



Easy-Load pump head




4 prong hypodermic

tubing diffuser

CEE shop

1/4” plug Cole Parmer H-06372-50 1/4” union Cole Parmer H-06372-50

stainless steel hypodermic

tubing

McMaster Carr

gas diffusing stone

Fisher Scientific 11-139B

99


1) Prepare the Ps. putida culture starting 48 hours before lab.

2) Prepare 100 mL glucose stock solution.

3) Attach one Easy-Load pump head to the pump drives and plumb with size 18 tubing connected to the hypodermic diffuser.

4) Verify that DO probes are operational, stable, and can be calibrated.

5) Mount DO probes on magnetic stirrers. (Use large stirbars.)

6) Use 100 mL plastic beakers containing 50 mL of bacteria suspension. The open tops will result in negligible oxygen transfer during the course of the experiments.

7) Prepare 1 L of deoxygenated distilled water right before class using the techniques outlined in the gas transfer lab (see page 154).

Glucose Stock Solution

C6H12O6 + 6O2 ⇒ 6CO2 + 6H2O

210gOL

6 12 6 6 12 62 26 12 6

2 2 6 12 6

1 180100.1 L = 0.9375 g C H O in 100 mL

32 6moleC H O gC H OgO moleO

L gO moleO moleC H O⋅ ⋅ ⋅ ⋅

Glucose Dilutions

2

10100 100

10000mgBOD L

mL LL m g O

µ⋅ ⋅ =

100 µL in 100 mL will provide 10 mg/L BOD 10 µL of stock solution diluted into 100 mL provides 1 mg BOD/L.

Table 4. BOD dilution water stock solutions. Use 10 mL per liter of each of the 4 solutions to prepare 10x BOD dilution water.

phosphate buffer M.W. g/L mg/100

mL µM

KH2PO4 136.09 8.5 850 62.46 K2HPO4 174.18 21.7

5 2175 124.87

Na2HPO4 · 7H2O 268.07 33.4 3340 124.60 NH4Cl 53.49 1.7 170 31.78

Magnesium

sulfate

MgSO4 120.39 11 1100 91.37

Calcium chloride CaCl2 110.99 27.5 2750 247.77

Ferric chloride

FeCl3 · 6H2O 270.3 0.25 25 0.925

100


Methane Production from Municipal Solid Waste

Introduction Archaeological investigations of landfills have revealed that biodegradable wastes

can be found — virtually intact — 25 years after burial. We know that landfills contain bacteria with the metabolic capability to degrade many of the materials that are common components of municipal refuse. The persistence for decades of degradable materials in the presence of such organisms appears somewhat paradoxical. In this experiment students will explore the factors that influence biodegradation of waste materials in landfills. Although recycling has significantly reduced the amount of landfill space dedicated to paper and other lignocellulosics, paper products are still a significant fraction of the solid waste stream. In this laboratory students will measure the rate and extent of anaerobic degradation of newsprint, Kraft paper, coated paper, and food scraps.

Theory Over 150 million tons of municipal solid waste (MSW) are generated every year in

the United States, and more than 70% of the MSW is deposited in landfills (Gurijala and Suflita 1993). Paper constitutes the major weight fraction of MSW, and this laboratory will focus on the biodegradation of that component. Anaerobic biodegradation of paper produces methane and carbon dioxide. Methane is a fuel and is the major component of natural gas. Methane produced in sanitary landfills represents a usable but underutilized source of energy. Energy recovery projects are frequently rejected because the onset of methane production is unpredictable and methane yields vary from 1-30% of potential yields based on refuse biodegradability data (Barlaz, Ham et al. 1992). The low methane yields are the result of several factors that conspire to inhibit anaerobic biodegradation including low moisture levels, resistance to biodegradation, conditions that favor bacterial degradation pathways that do not result in methane as an end product, and poor contact between bacteria and the organic matter.

Characteristics of municipal solid waste

The physical composition of residential municipal solid waste (MSW) in the United States is given in Table 1. The fractional

Table 1. Typical physical composition of residential MSW in 1990 excluding recycled materials and food wastes discharged with wastewater (Tchobanoglous, Theisen et al. 1993)

Component Range Typical Organic (% by weight) (% by weight)

food wastes 6-18 9.0 paper 25-40 34.0

cardboard 3-10 6.0 plastics 4-10 7.0 textiles 0-4 2.0 rubber 0-2 0.5 leather 0-2 0.5

yard wastes 5-20 18.5 wood 1-4 2.0

Organic total 79.5 Inorganic

glass 4-12 8.0 tin cans 2-8 6.0

aluminum 0-1 0.5 other metal 1-4 3.0

dirt, ash, etc. 0-6 3.0 Inorganic total 20.5

101


contribution of the listed categories has evolved over time, with a trend toward a decrease in food wastes because of increased use of kitchen food waste grinders, an increase in plastics through the growth of their use for packaging, and an increase in yard wastes as burning has ceased to be allowed by most communities (Tchobanoglous, Theisen et al. 1993). Excluding plastic, rubber, and leather, the organic components listed in Table 1 are, given sufficient time, biodegradable.

Although recycling efforts divert a

significant fraction of paper away from landfills, paper continues to be a major component of landfilled waste. The types of paper found in MSW are listed in Table 2.

The elemental composition of

newsprint and office paper are listed in Table 3.

The major elements in paper are

carbon, hydrogen, and oxygen that together constitute 93.5% of the total solids. The approximate molecular ratios for newspaper and office paper are C6H9O4 and C6H9.5O4.5 respectively.

Biodegradation of cellulose, hemicellulose, and lignin

Cellulose and hemicellulose are the principal biodegradable constituents of refuse accounting for 91% of the total methane potential. Cellulose forms the structural fiber of many plants. Mammals, including humans, lack the enzymes to degrade cellulose. However, bacteria that can break cellulose down into its subunits are widely distributed in natural systems, and ruminants, such as

Table 2. Percentage distribution by weight of paper types in MSW (Tchobanoglous, Theisen et al. 1993)

Type of paper Range Typical newspaper 10-20 17.7 books and magazines

5-10 8.7

commercial printing

4-8 6.4

office paper 8-12 10.1 other

paperboard 8-12 10.1

paper packaging 6-10 7.8 other

nonpackaging paper

8-12 10.6

tissue paper and towels

4-8 5.9

corrugated materials

20-25 22.7

Total 100.0

Table 3. Elemental composition of two paper types on a dry weight basis (Tchobanoglous, Theisen et al. 1993).

Constituent Newsprint Office Paper C 49.1% 43.4% H 6.1% 5.8% O 43.0% 44.3%

NH4-N 4 ppm 61 ppm NO3-N 4 ppm 218 ppm

P 44 ppm 295 ppm PO4-P 20 ppm 164 ppm

K 0.35% 0.29% SO4-S 159 ppm 324 ppm

Ca 0.01% 0.10% Mg 0.02% 0.04% Na 0.74% 1.05% B 14 ppm 28 ppm Zn 22 ppm 177 ppm Mn 49 ppm 15 ppm Fe 57 ppm 396 ppm Cu 12 ppm 14 ppm

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cows, have these microorganisms in their digestive tract. Cellulose is a polysaccharide that is composed of glucose subunits (see Figure 1).

Another component of the walls of plants is hemicellulose, which sounds similar to cellulose but is unrelated other that that it is another type of polysaccharide. Hemicelluloses made up of five carbon sugars (primarily xylose) are the most abundant in nature.

Lignin is an important structural component in plant materials and constitutes roughly 30% of wood. Significant components of lignin include coniferyl alcohol and syringyl alcohol subunits (Figure 2).

The exact chemical structure of lignin is not known but its reactivity, breakdown products, and the results of spectroscopic studies reveal it to be a polymeric material containing aromatic rings with methoxy groups (-OCH3) (Tchobanoglous, Theisen et al. 1993). One of the many proposed structures for lignin is shown in Figure 3.

Degradation of lignin requires the presence of moisture and oxygen and is carried out by filamentous fungi (Prescot, Harley et al. 1993). The biodegradability of lignocellulosic materials can be increased by an array of physical/chemical processes including pretreatment to increase surface area (size reduction), heat treatment, and treatment with acids or bases. Such treatments are useful when wood and plant materials are to be anaerobically degraded to produce methane. Research on this topic has been performed by Cornell Prof. James Gossett (Gossett and McCarty 1976; Chandler, Jewell et al. 1980; Gossett, Stuckey et al. 1982;

Figure 1. Cellulose (two glucose subunits are shown).

-C-C-C-HO-

CH O3

CH O3

-C-C-C-HO-

CH O3

Figure 2. Coniferyl (left) and syringyl (right) subunits of lignin.

CH O3

C

OCH3-OHC

H

OC

H

H

H

CH

OCH3

-OH

C H

C

H

H

O

C

HCH O

3

O

C H

H

C

O

H

H

CH

OH

OCH3

-OH

OCH3

C

H

C

H

CH

O

O

C

CHH

HO

H

CHHO C

H H CH

O

-OH

OCH3

H

HO

OCH3

C

O

C

CH O

H

OH

OCH3

C H

C O

OCH3

C

CHHO

H

OHH

OC

COH

H

HH

C H

C H

C

H

HH

OOH

O

Figure 3. A postulated formulation for spruce lignin (by (Brauns 1962), as cited by (Pearl 1967)). This structure is suggested by spectroscopic studies and the chemical reactions of lignin.

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Pavlostathis and Gossett 1985a; Pavlostathis and Gossett 1985b). Three major groups of bacteria are involved in the conversion of cellulosic material

to methane (Zehnder 1978): (1) the hydrolytic and fermentative bacteria that break down biological polymers such as cellulose and hemicellulose to sugars that are then fermented to carboxylic acids, alcohols, carbon dioxide and hydrogen gas, (2) the obligate hydrogen reducing acetogenic bacteria that convert carboxylic acids and alcohols to acetate and hydrogen, and (3) the methanogenic bacteria that convert primarily acetate and hydrogen plus carbon dioxide to methane. Sulfate reducing bacteria (SRB) may also play a role in the anaerobic mineralization of cellulosic material. In the presence of sulfate, the degradation process may be directed towards sulfate reduction by SRB with the production of hydrogen sulfide and carbon dioxide (Barlaz, Ham et al. 1992).

Cellular requirements for growth The availability of oxygen is a prime determinant in the type of microbial

metabolism that will occur. Microbial respiration of organic carbon is a combustion process, in which the carbon is oxidized (i.e., is the electron donor) in tandem with the reduction of an electron acceptor. The energy available to microorganisms is greatest when oxygen is used as the electron acceptor and therefore aerobic metabolic processes will dominate when oxygen is available. Some microorganisms require oxygen to obtain their energy and are termed “obligate aerobes.” In the absence of oxygen, other electron acceptors such as nitrate (NO3

-), sulfate (SO4-2) and carbon

dioxide (CO2) can by used. Organisms that can only exist in an environment that contains no oxygen are termed “obligate anaerobes.” Organisms that have the ability to grow in both the presence and the absence of oxygen are said to be “facultative.”

The availability of nutrients can limit the ability of cells to grow and consequently the extent of biodegradation. Nitrogen and/or phosphorous constitute important nutrients required for cell synthesis. Inorganic bacterial nutritional requirements also include sulfur, potassium, magnesium, calcium, iron, sodium and chloride. In addition, inorganic nutrients needed in small amounts (minor or trace nutrients) include zinc, manganese, molybdenum, selenium, cobalt, copper, nickel, vanadium and tungsten. Organic nutrients (termed “growth factors”) are also sometimes needed (depending on the microorganism) and include certain amino acids, and vitamins (Metcalf & Eddy 1991).

Environmental conditions such as pH, temperature, moisture content, and salt concentration can have a great influence on the ability of bacteria to grow and survive. Most bacteria grow in the pH range from 4.0 to 9.5 (although some organisms can tolerate more extreme pH values), and typically grow best in the relatively narrow range from 6.5 to 7.5 (Metcalf & Eddy, 1991). Microorganisms have a temperature range over which they function best, and are loosely characterized as phychrophilic (ability to grow at 0°C), mesophilic (optimal growth at 25-40°C) or thermophilic (optimal growth above 45-50°C) (Brock 1970). Many common methanogens are mesophilic. Elevated temperatures also favor faster reaction rates.

While some microorganisms are very tolerant of low moisture conditions, active microbial growth and degradation of organic matter necessitates that water not be a scarce resource. Cells take water in through their semi-permeable membrane surface

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by osmosis. This uptake mechanism requires that the solute concentration inside the cell be higher than that of the outside media. Organisms that grow in dilute solutions can not tolerate high salt concentrations because their normal osmotic gradient is reversed and they can not take in water. Some cell strains, termed “halophiles” are adapted for growth at very high salt concentrations.

The above factors suggest that bacterial degradation of MSW to produce methane will occur optimally at circumneutral pH, low ionic strength, in the absence of oxygen, nitrate and sulfate, in the presence of moisture and nutrients, and under mesophilic conditions.

Estimates of paper biodegradability Volatile solids (VS) content (determined by weight loss on ignition at 550°C) has

been used to estimate the biodegradability of MSW components, but this measure overestimates the biodegradability of paper. Paper products have a very high volatile solids content. Newsprint, office paper, and cardboard have VS of 94%, 96.4%, and 94% respectively (Tchobanoglous, Theisen et al. 1993). Paper products also can have a high content of lignocellulosic components that are only slowly degradable. Lignin constitutes approximately 21.9%, 0.4% and 12.9% respectively of the VS in newsprint, office paper, and cardboard. Lignin content and biodegradability are strongly correlated and thus lignin content can be used to estimate biodegradability and potential methane production. Chandler et al. (1980) found a relationship between lignin content and biodegradable volatile solids using a wide variety of waste materials. The empirical relationship suggests that not only is lignin not easily biodegraded, but that lignin also reduces the biodegradability of the nonlignin components. This reduction in biodegradability may be caused by lignin polymeric material physically preventing enzymatic access to the nonlignin components. The relationship is

biodegradable ligninVS 2.8VS 0.83= − + 9.1

where VSbiodegradable is the biodegradable fraction of the volatile solids and VSlignin is the fraction of volatile solids that are lignin. From equation 9.1 the maximum destruction of VS is limited to about 83%, a limitation due to the production of bacterial by-products. The high concentration of lignin in newsprint makes it much less biodegradable than more highly processed office paper (Table 4).

Energy recovery from MSW Energy could be recovered from

MSW by direct combustion in an incinerator or by anaerobic biodegradation and production of methane. Proximate analysis is used to measure moisture content, volatile matter, fixed carbon (combustible but not volatile), and ash. Proximate analysis can be used to predict ash

Table 4. Biodegradability of selected components of MSW (Tchobanoglous, Theisen et al. 1993)

VS/TS Lignin/VS VSbiodegradable* Type of waste % % %

mixed food 7-15 0.4 82 newsprint 94 21.9 22

office paper 96.4 0.4 82 cardboard 94.0 12.9 47

* Obtained by using equation 9.1

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production from incineration. The energy content is measured in a bomb calorimeter. Proximate analysis results and energy content of MSW are given in Table 5.

Gas production

from anaerobic digestion is typically 30% CO2 and 70% CH4. The methane is a valuable fuel and has an energy content of 802.3 kJ/mol or 50 MJ/kg. The combustion of methane produces only carbon dioxide and water.

Because paper products are a major fraction of MSW and paper energy content is significant, the majority of energy in MSW is contained in paper products. Incineration or methane production can be used to capture some of this available energy.

4 2 2 2CH +2O CO +2H O→ 9.2

Effect of MSW particle size The large size of pieces of MSW is suspected to decrease the ability of microbes to

degrade the material. Landfill gas production has been correlated with refuse particle size (Ferguson 1993). The effect of particle size reduction was initially explained by the resultant increase in surface area available for microbial attach. Laboratory studies under saturated conditions, however, suggest that size reduction, even down to a few microns or tens of microns has little effect on the rate of degradation. According to Ferguson (1993), surface area increases only slightly with decreasing particle size for platey and fibrous particles such as paper. Thus the effect of size reduction on the methane production in landfills may be that relatively large pieces of plastic, paper, or other material shield the materials beneath them from infiltrating water. The shielded material may remain too dry for biodegradation. Pulverization breaks down the impermeable barriers and more of the waste is exposed to water (Ferguson 1993).

Potential methane production from municipal solid waste Under anaerobic conditions microorganisms can produce both CO2 and CH4

(methane) without consuming any oxygen. Other significant end products include odorous gases such as ammonia (NH3), and hydrogen sulfide (H2S) (see Figure 4). Because anaerobic biodegradation produces gas it is possible to monitor the extent and rate of anaerobic biodegradation by measuring gas production (Suflita and Concannon 1995).

Table 5. Proximate analysis and energy content of selected components of MSW (Tchobanoglous, Theisen et al. 1993).

moisture volatile matter

fixed carbon

ash energy as

collected

energy dry

Type of waste % % % % (MJ/kg) (MJ/kg) fats 2 95.3 2.5 0.2 37.5 38.3

mixed food 70 21.4 3.6 5 4.2 13.9 fruit waste 78.7 16.6 4 0.7 4.0 18.6 meat waste 38.8 56.4 1.8 3.1 17.7 29.0 cardboard 5.2 77.5 123 5 16.4 17.3 magazines 4.1 66.4 7 22.5 12.2 12.7 newsprint 6 81.1 11.5 1.4 18.6 19.7

mixed paper 10.2 75.9 8.4 5.4 15.8 17.6 waxed cartons 3.4 90.9 4.5 1.2 26.3 27.3

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Gas production Because anaerobes get relatively

little energy from the organic matter their conversion of carbon to cell material (synthesis) is much lower than for aerobes. Typically 10% of the organic matter may be converted to anaerobe cell mass. Thus the majority of the biodegraded organic matter is converted to gas and the gas production can be used as a measure of biodegradation. The ideal gas law is used to determine the moles of gas produced from the pressure, volume, and temperature.

PV

nRT

= 9.3

The pressure in the sealed test bottles that will be used in this laboratory is initially atmospheric. Because the number of moles is a linear function of the pressure we can write

PV

nRT

∆∆ = 9.4?

where ?P is the change in pressure relative to the initial pressure in the bottle. In these experiments the bottle volume is 120 mL and the maximum recommended

pressure increase is 80 kPa (12 psi). The volume of liquid in the bottles is 20 mL and the volume contributed by solids is expected to be negligible. Thus the nominal volume of gas in the bottles will be 100 mL. Solving for the number of moles of gas (CH4 and CO2) produced by anaerobic digestion

( )( )

( )

3 6 3

3

80 10 100 103.13 mmole C

8.31 308

x Pa x mn

Pa mK

mol K

−

∆ = = ⋅ ⋅

9.5

The molecular formula of cellulose is C6H10O5 and thus 27 g of cellulose has 1 mole of carbon. The relation obtained in equation 9.5 is used to determine the maximum amount of cellulose that can be anaerobicly degraded without exceeding 80 kPa in the bottles.

27 mg cellulose

3.13 mmole C 84 mg cellulosemmole C

⋅ = 9.6

The mass of paper containing 84 mg of biodegradable cellulose can be obtained using Table 4 and the results of equation 9.6. The mass of dry newspaper that will produce a pressure increase of 80 kPa is

Organic Matter

Nutrients CH4 CO2NH3 H2S

Refractory organic matterHeat

CellsH2O New Cells

Figure 4. Reactants and products for anaerobic degradation of organic matter.

107


biodegradable

biodegradable

84 mg VS VS TS 400 mg dry newspaper80 kPa 0.22 VS 0.94 VS 80 kPa

⋅ ⋅ = 9.7

Similar calculations can be performed for other types of waste. The maximum mass of glucose (CH2O has 30 g of glucose per mole of carbon) is

30 mg glucose

3.13 mmole C 94 mg glucosemmole C

⋅ = 9.8

Although glucose is expected to be completely biodegradable, a small amount of glucose will be converted into refractory cell byproducts.

The above calculations are based on the assumption that all of the gas produced is volatile and is not dissolved. Carbon dioxide is soluble and thus some of the CO2 produced will be dissolved and will not result in increased pressure.

Acid neutralizing capacity requirements The high partial pressure of CO2 resulting from anaerobic biodegradation causes a

high concentration of carbonic acid *2 3H CO and thus would result in a reduced pH

if there were insufficient Acid Neutralizing Capacity (ANC). The amount of ANC required to counteract the high partial pressure of CO2 can be obtained from the Henry’s constant for dissolution of CO2, and from the dissociation constant for carbonic acid.

2

*2 3

HCO

H COK

P

= 9.9

where KH has a value of 3.12 x 10-4 moles/J. The first dissociation constant for carbonic acid is

3

1 *2 3

H HCOK

H CO

+ − =

9.10

where K1 has a value of 10-6.3. The definition of ANC for a carbonate system in equilibrium with the gas phase is

( )2

1 20

2CO H wP K K

ANC HH

α αα

++

= + + − 9.11

Where α0, α1, α2 are the fractions of total carbonate present as carbonic acid *

2 3H CO , bicarbonate 3HCO − , and carbonate 23CO− respectively and Kw is the

dissociation constant for water. At circumneutral pH the hydrogen ion, hydroxide ion, and carbonate ion concentrations are negligible and equation 9.11 simplifies to

2 1

0

CO HP KANC

α

α= 9.12

108


The ratio of bicarbonate to carbonic acid may be determined from equation 9.10. Solving for the ratio of bicarbonate to carbonic acid:

3 1 1

*02 3

HCO K

H CO H

αα

−

+

= =

9.13

Equation 9.13 can be substituted into equation 9.12 to obtain

2 1CO HP K KANC

H +=

9.14

An estimate of the ANC required to maintain a neutral pH under a pressure of 30 kPa of CO2 can be obtained by substituting appropriate values into equation 9.14.

( ) ( )4 4 6.3

7

3 10 3.12 10 10

1047 /

molesx Pa x M

N mANC

MANC meq L

− −

−

⋅ ≅

≅

9.15

The basal medium that will be used in this laboratory contains 71 meq/L ANC from sodium bicarbonate. If the pressure of CO2 reaches 60 kPa (30 kPa initial pressure plus 30 kPa from the production of CO2 during an experiment) then solving equation 9.14 for pH shows that (given the 71 meq ANC in the basal medium and a CO2 pressure of 60 kPa) the pH will drop to 6.88. Thus, the basal medium is sufficiently buffered to protect against significant pH changes.

Carbon dioxide solubility At pH less than ˜9 the inorganic carbon will partition into three species, gaseous

2CO , aqueous *2 3H CO , and aqueous 3HCO− .

*2 2( ) ( ) 2 3 3total gCO CO H C O HCOn n n n −= + + 9.16

The number of moles of the inorganic carbon species can be determined based on the partial pressure of 2CO , the ANC of the liquid and the gas and liquid volumes. The moles of gaseous 2CO is obtained from the ideal gas law

2

2( )g

CO gCO

P Vn

RT= 9.17

The number of moles of *2 3H CO is obtained from the Henry’s law constant.

* 22 3CO H lH C O

n P K V= 9.18

The concentration of bicarbonate, 3HCO− , is equal to the ANC (for pH < 9).

3

lHCOn ANC V− = ⋅ 9.19

109


The total number of moles of inorganic carbon is the sum of the three species.

( )2

2 2( )total

CO gCO CO H l

P Vn P K ANC V

RT= + + 9.20

Therefore, the number of moles of inorganic carbon in an enclosed volume is a linear function of the partial pressure of CO2 (Figure 5).

The pH will change as the partial pressure of CO2 changes as shown in equation 9.14. Solving for the concentration of hydrogen ions equation 9.14 becomes

2 1CO HP K KH

ANC+ = 9.21

The relationship between pH and partial pressure of CO2 is shown in Figure 5.

The basal medium to be used in this experiment will be purged with a 30:70 mixture of carbon dioxide and nitrogen prior to use. As shown in Figure 5 the pH of the basal medium is expected to rise to approximately 7.17. The headspace will also be purged with the same gas mixture and thus there will be 5.6 mmoles of inorganic carbon in the bottles initially. After the anaerobic biodegradation has gone to completion, the carbon dioxide concentration will be measured by gas chromatography and the gas pressure by pressure sensors and thus the partial pressure of carbon dioxide will be known. Figure 5 or equation 9.20 can be used to determine the final mass of inorganic carbon in the bottles. The difference between the initial and final inorganic carbon concentration can be used to determine the amount of organic carbon converted to carbon dioxide.

Methane solubility The Henry’s constant for methane ( ( )4H C HK ) at 25°C is 1.48 x 10-5 mol/J (Mackay

and Shiu 1981). Methane is significantly less soluble than carbon dioxide and does not form other soluble aqueous species. The mass of gaseous and dissolved methane is given by equations 9.22 and 9.23.

6.5

6.6

6.7

6.8

6.9

7.0

7.1

7.2

7.3

7.4

7.5

0 20 40 60 80 100

Partial pressure of CO2 (kPa)

4.0

4.5

5.0

5.5

6.0

6.5

7.0

7.5

8.0

8.5

9.0

inor

gani

c ca

rbon

(mm

oles

)

pH

mmolesinorganic carbon

Figure 5. Number of moles of inorganic carbon and pH as a function of the partial pressure of carbon dioxide given gas and liquid volumes of 60 mL and ANC of 71 mmoles/L.

110


4

4 ( )CH g

CH g

P Vn

RT= 9.22

( )4 4 4( )CH aq CH lH C Hn P K V= 9.23

The ratio of the mass of gaseous to dissolved methane gives an indication of the significance of dissolved methane.

( )

4

4 4

( )

( )

CH g g

CH aq l H C H

n V

n V K RT= 9.24

Substituting appropriate values into equation 9.24

( )

4

4

( )

5( ) 1.48 10 8.31 308

CH g g

CH aql

n V

mol JnV x K

J mol K−

= ° °

9.25

If the gas and liquid volumes are approximately equal there will be approximately 26 times as much methane in the gaseous phase as in the dissolved phase. This ratio is independent of the methane partial pressure. The total number of moles of methane can be obtained from the partial pressure of methane in the gaseous phase.

( )4 4 4( )g

CH total CH lH CH

Vn P K V

RT

= +

9.26

The partial pressure of methane will be determined from the pressure in the bottle and mass of methane as measured by the gas chromatograph.

Temperature effects The temperature of the bottles directly affects the pressure of gas as well as

influences the rate of gas production by the microbes. Cummings and Stewart found that methane production was sharply inhibited by temperatures in excess of the optimum (37°C) and was undetectable at 20°C (1995). However, Suflita and Concannon (1995) reported anaerobic digestion at “room temperature” over a period of 2 months. If desired a constant temperature water bath can be used to keep all of the digesters at a constant and optimal temperature (35°C) for anaerobic degradation.

Experiment description The experimental setup is a flexible system for obtaining data on the anaerobic

decomposition of various organic materials by measuring the pressure of the gas produced. A schematic of the experimental setup is shown in Figure 6.

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Bacterial degradation of

selected materials will be assayed by placing known quantities in 120 mL bottles, inoculating with an active anaerobic mixed culture, sealing the bottles with rubber septa and aluminum crimp caps, and monitoring gas pressure and composition over time. Anaerobic digester supernatant from the Ithaca Wastewater Treatment Plant will be used as a source of microbes. The bottles will be monitored for biogas production with pressure sensors connected with a needle through the septa (see Figure 6). Gas composition will be determined by periodic analysis of methane (CH4) and carbon dioxide (CO2) via gas chromatography with a thermal-conductivity detector.

Gas production measurements will be automated by using pressure sensors in a procedure comparable to that described by Suflita and Concannon (1995). With this technique, a large number of bottles can be monitored with automated data acquisition by a single computer, allowing a wide variety of chemical and environmental parameters to be explored. The automated acquisition of gas data is necessary due to the numbers of bottles and length of incubation (ca. 4 weeks) anticipated. This experiment will be set-up and left virtually unattended while other laboratory exercises continue in intervening weeks.

Each sample type should be cut into small enough pieces to easily insert into the bottle. Students may also be interested in exploring biodegradation of other organic components of municipal solid waste (banana peels, rags, plastic bags, etc.). Table 6 suggests one configuration of several sample types. Each sample will receive 15 mL of basal medium.

One control should be “unamended” (i.e., with no waste) and contain the microbial inoculum and the O2-free water to monitor any gas production attributable just to the added sludge, one “positive”

Hypodermic needle

Crimp cap with septa

Pressure sensor

Anaerobic solution

analogto

digital Power Supply (12 V)

Connector panel

Multiplexer

RJ 11 plug

Serum bottle

35º C incubator

Hypodermic needle

Crimp cap with septa

Pressure sensor

Anaerobic solution

analogto

digital Power Supply (12 V)

Connector panel

Multiplexer

RJ 11 plug

Serum bottle

35º C incubator

Figure 6. Experimental setup (not to scale). Pressure generated by microbial gas production is monitored by pressure sensors.

Table 6. Suggested sample preparation. Each group does 2 sample types with 2 replicates. Each section does 2 water and 2 inoculum controls.

Sample Size

Inoculum Replicates

Sample Type mg mL # environmental

control none 0 2

bacterial control none 5 2 positive control

(glucose) 90 5 2

filter paper 50 5 2 cardboard ? 5 2

office paper ? 5 2 newsprint ? 5 2

Student selected organic materials

? 5 up to 4 vials

112


control should be the microbial inoculum, plus 90 milligrams of glucose and the O2-free water (to verify that the microbial population is active in the added sludge), and one control should be plain O2-free water to control for variations in temperature, and air pressure. The sample sizes of the various samples should be determined so that the bottles will not generate pressure greater than 80 kPa.

Experimental methods

Safety concerns

1) Municipal wastewater sludge will be used as a source of microbes. The sludge may contain biological and/or chemical hazards and should be handled accordingly.

2) Biological production of gas will generate pressure in a closed container. Testing has shown that this system is safe up to at least 200 kPa (30 psi). At approximately this pressure the needle is typically forced out of the septa. If the bottle is not vented the pressure can increase until the crimp cap is forced off. Bottles should not be capped for very long before the needles are inserted and pressure monitoring begins. The pressure trends should be monitored and, if excessive pressures are produced, the bottles must be vented and/or the temperature of the bath may be reduced.

3) Sharp needles are used in the experimental setup and precautions should be taken to avoid puncturing unintended objects (including students).

Analysis of moisture content and volatile solids The fraction of volatile solids in the paper samples is the maximum that could

possibly be degraded. Note that paper products cannot be ashed accurately because the strong flames easily carry some of the ashes away. Steps for sample moisture content and volatile solids fraction follow: 1) Weigh an aluminum boat 2) Weigh the aluminum boat with an organic sample (boat + water + VS + ash) 3) Dry in the 105°C oven 4) Cool in a desiccator 5) Weigh (boat + VS + ash) 6) Ash in the 550°C muffle furnace 7) Cool in a desiccator 8) Weigh (boat + ash)

Sample preparation The bottles will be purged initially with a mixture of 30% CO2 and 70% N2 to

remove any O2 and to establish an initial carbon dioxide concentration so that the initial pH is not excessively high. If no carbon dioxide were present in the purging

113


gases the carbonic acid would be stripped out of solution and the pH would rise in the basal medium.

Acquisition of pressure data The biogas pressure will be measured indirectly by the pressure sensors. The

specified sensors work from zero to 100 kPa (6.89476 kPa/psi). They will withstand 2.5 times rated pressure (i.e., 250 kPa) but their output may be erroneous above the upper limit of the working range. The output of the pressure sensors is zero to 0.100 volts with 0.100 volts indicating approximately 100 kPa. The outputs of the sensors are fed through a 32-channel multiplexer/signal conditioner, an A/D converter board and are monitored using LabVIEW software.

Gas chromatograph analysis for separation of CO2, CH4 and N2 (Optional)

Permanent gases can be analyzed using a thermal conductivity detector (TCD) on a gas chromatograph. The thermal conductivity detector measures the rate at which heat is transported from the detector. If a gas with a different thermal conductivity than the carrier gas passes through the detector a peak is detected. A flame ionization detector could be used to measure methane, but would not be able to detect carbon dioxide or nitrogen since they do not burn. A micropacked column containing packing designed for analyses of permanent gases and light hydrocarbons (Supelco Carboxen 1004) is used to separate the gases.

The TCD must be calibrated with known masses of the gases of interest. Nitrogen, carbon dioxide and methane are available as compressed gases and can be sampled at atmospheric pressure by opening a valve in a compressed gas line slightly and sampling the discharge with a gas tight syringe. The ideal gas law is used to calculate the moles of gas. The current atmospheric pressure in Ithaca is available through the World Wide Web at http://cuinfo.cornell.edu/Ithaca/Weather/. If the atmospheric pressure is reported in inches of mercury it can be converted to Pascals by multiplying by 3386 Pascals/Inch of Hg. The temperature of the laboratory is available from the pH meters equipped with temperature probes. If a 100 µL gas sample is used, the atmospheric pressure is 100 kPa, and the temperature is 22°C then the number of moles of gas are calculated as:

( ) ( )

( )

-9 3

3

100,000 Pa 100x10 m4.08 mol

8.31 295n

Pa mK

mol K

µ= = ⋅ ⋅

9.27

The number of moles of gas is independent of the type of gas. The relationship between peak area and moles of gas is calculated by analyzing a known number of moles of each gas. The TCD response will be different for each gas since the thermal conductivity of each gas is different.

Experimental method (short version)

1) Dry 2 – 2 g samples for each sample type in the 105°C oven. 2) Take dried organic sample from the oven.

114


3) Keep 1 dried sample and determine the VS of the other sample. 4) Weigh appropriate amounts of the various dried samples for methane production. 5) Load bottles with organic samples (cut to smaller size as needed). 6) Add 15 mL of basal medium to each of the bottles. 7) Add 5 mL of inoculum to each of the bottles. 8) Purge the headspace of the bottles with an oxygen-free gas stream that is 30%

CO2 and 70% N2.

9) Seal the bottles. 10) Insert the pressure sensor hypodermic needle into the bottle. 11) Sample the bottle pressures using the data acquisition software (take samples

every hour and save the data as \\Enviro\enviro\Courses\453\methane\pressure).

Gas Analysis Method

1) Calibrate the gas chromatograph using methane and carbon dioxide and using 20 µL samples

2) Take an initial headspace gas sample and analyze it using the gas chromatograph. 3) Sample gas composition after gas production has ceased using the gas

chromatograph.

Prelab questions 1) Estimate the mass of cardboard and the mass of office paper that will produce a

pressure rise of 80 kPa in the sample bottles at 35°C if the headspace volume is 100 mL. Use the predicted biodegradability based on the lignin content of the paper.

Data analysis Perform the analysis on the data from your lab section.

1) Calculate total gas production in moles. For each sample use the record of pressure vs. time to determine if the reaction appears to have gone to completion.

2) For your samples, compare volatile solids (VS) and gas production by converting the mass of volatile solids to moles of carbon using an approximate molecular formula for the sample. The molecular formula for the volatile fraction of paper can be approximated by C6H10O5. Calculate and plot the fraction of VS degraded as a function of time for each sample.

3) Compare the fuel value of the methane produced with the fuel value of the original sample for each of the samples. Use the estimates of the original fuel value (Table 5) and the measured methane production. The fuel value of glucose is 424.7 KJ/mole C. If you don’t have the gas composition of your samples, then assume 70% of the gas produced was methane.

115


Optional Analysis Requiring Gas Composition

1) Calculate the moles of CO2 and CH4 produced by your samples based on the gas chromatograph analysis. Include the effect of carbon dioxide solubility. Use the basal medium control to subtract the initial headspace as well as any gas production by the inoculum.

2) Calculate the final pressure based on the GC measurements and compare with the pressure transducer measurements. Remember that the pressure transducer measured gage pressure.

References Barlaz, M. A., R. K. Ham, et al. (1992). “Microbial chemical and methane production

characteristics of anaerobically decomposed refuse with and without leachate recycling.” Waste Management & Research 10(3): 257-267.

Brauns, F. E. (1962). “Soluble native lignin, milled wood lignin, synthetic lignin and the structure of lignin.” Holzforschung 16: 97-102.

Brock, T. D. (1970). Biology of Microorganisms. London, Prentice-Hall. Chandler, J. A., W. J. Jewell, et al. (1980). Predicting Methane Fermentation

Biodegradability. Biotechnology and Bioengineering Symposium No. 10, John Wiley & Sons, Inc.

Cummings, S. P. and C. S. Stewart (1995). “Methanogenic interactions in model landfill co-cultures with paper as the carbon source.” Letters in Applied Microbiology 20(5): 286-289.

DiStefano, T. D. (1992). Biological dechlorination of tetrachloroethene under anaerobic conditions, Cornell University.

Ferguson, C. C. (1993). “A hydraulic model for estimating specific surface area in landfill.” Waste Management & Research 11(3): 227-248.

Gossett, J. M. and P. L. McCarty (1976). “Heat Treatment of Refuse for Increasing Anaerobic Biodegradability.” AIChE Symposium Series 158(72): 64-71.

Gossett, J. M., D. C. Stuckey, et al. (1982). “Heat Treatment and Anaerobic Digestion of Refuse.” Journal of the Environmental Engineering Division 108: 437-454.

Gurijala, K. R. and J. M. Suflita (1993). “Environmental factors influencing methanogenesis from refuse in landfill samples.” Environmental Science and Technology 27(6): 1176-1181.

Mackay, D. and W. Y. Shiu (1981). “A critical review of Henry's law constants for chemicals of environmental interest.” Journal of Physical Chemical Reference Data 10(4): 1175-1199.

Metcalf & Eddy, I. (1991). Wastewater Engineering. Treatment, Disposal, and Reuse. New York, McGraw-Hill, Inc.

Pavlostathis, S. G. and J. M. Gossett (1985). “Alkaline Treatment of Wheat Straw for Increasing Anaerobic Biodegradability.” Biotechnology and Bioengineering 27: 334-344.

116


Pavlostathis, S. G. and J. M. Gossett (1985). “Modeling Alkali Consumption and Digestibility Improvement From Alkaline Treatment of Wheat Straw.” Biotechnology and Bioengineering 27: 345-354.

Pearl, I. A. (1967). The Chemistry of Lignin. New York, NY, Marcel Dekker, Inc. Prescot, L. M., J. P. Harley, et al. (1993). Microbiology. Dubuque, IA, Wm. C.

Brown, Publ. Suflita, J. M. and F. Concannon (1995). “Screening tests for assessing the anaerobic

biodegradation of pollutant chemicals in subsurface environments.” Journal of Microbiological Methods 21: 267-281.

Tchobanoglous, G., H. Theisen, et al. (1993). Integrated Solid Waste Management. Engineering Principles and Management Issues. New York, McGraw-Hill, Inc.

Zehnder, A. J. B. (1978). Ecology of methane formation. Water Pollution Microbiology. R. Mitchell. New York, John Wiley & Sons. 2: 349-376.

117


Lab Prep Notes

Setup

1) Use anaerobic digester supernatant as inoculum source. Place supernatant under fume hood. Use 5 mL per sample.

2) Setup 10 port purger with CO2 and N2 gas metered through rotometers. The top ball should be at 24 mm for CO2 and at 84 mm for N2.

3) Set the GC with 300 Kpa column pressure, 180ºC oven, 250ºC injector and detectors, and 1.2 minute run time. Use 20 µL sample. The gases should come out in the order N2, CH4, and CO2 at 0.44, 0.72, and 1 minute respectively. (Only if you are doing the optional GC analysis)

4) 4 samples/group plus 2 inoculum blanks and 2 water blanks.

Class Plan

1) Sign up for samples 2) Each group chooses 2 types of

samples 3) Dry samples in oven 4) Ash 1 of the 2 samples


Description Vender Catalog 500 µl syringe w/

valve Supelco 2-2272

side port needle Supelco 2-2289 Carboxen 1004 micropacked

column

Supelco 1-2846

Hp 5890 Series II GC

Hewlett-Packard 5890A

TCD kit Hewlett-Packard 19232E 1/8" column

adapter Hewlett-Packard option 095

pressure regulators Hewlett-Packard L43 RS232C board Hewlett-Packard option 560

Helium Cornell Stores Wrist action Shaker Fisher Scientific 14-260

Vials Supelco 3-3111 Aluminum crimp

tops Fisher 03-375-23C

Butyl stopper Fisher 03-375-22AA Crimping tool Supelco 3-3280

EPDM stoppers-13x20 mm

Sigma Z16607-3

luer lock needles 21 gauge

Fisher 14-826-5B

Pressure transducer, 0 to 15

psig

Omega PX136-015GV

12 V DC Power supply

Omega PSS-12

Incubator Fisher 11-690-650D Multiplexer National

Instruments

4 slot chassis SCXI-1000

776570-01

32 channel SCXI-1100

776572-00

SCXI-1200 parallel port

776783-00

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Table 11. Gas chromatograph conditions

gas pressure flow

carrier (He) kPa 5 mL/min Ref 15 mL/min

temperatures °C oven (isothermal)

Injector 250 TCD 250

Column Supplier Catalog

number Carboxen 1004 micropacked

column

Supelco 1-2846

Table 8. Reagents

Reagent Vender Catalog basal medium NA

glucose Aldrich 15,896-8 paper (various

types) NA

Whatman Filter Paper (No. 1)

Fisher Scientific 09-805-1A

Table 9. Basal medium for anaerobic growth (DiStefano 1992).

Compound Quantity (per

liter) NH4Cl 200 mg

K2HPO4·3H2O 100 mg KH2PO4 55 mg

MgCl2·6H2O 200 mg Resazurin 1 mg

FeCl2·4H2O 100 mg Trace Metals

Solution 10 mL

Na2S·9H2O 500 mg NaHCO3 6 g

The first six compounds are added to distilled-deionized water, then purged with N2 until solution turns from blue to pink. The remaining components are added, followed by a 15-minute purge with the 70% N2/30% CO2 gas mixture.

Table 10. Trace metals for anaerobic growth (DiStefano 1992).

Compound Quantity (mg/L)

MnCL2·4H2O 100 CoCl2·6H2O 170

ZnCl2 100 CaCl2·2H2O 251

H3BO3 19 NiCl2·6H2O 50

Na2MoO4·2H2O 20

119

Volatile Organic Carbon Contaminated Site Assessment


Introduction Roughly 75 percent of the major cities in the U.S. depend, at least in part, on

groundwater for their water supply. Various estimates of the nationwide extent of groundwater contamination are stated to range from one to over two percent of the nation's usable groundwater (Council on Environmental Quality, 1981). Volatile organic compounds (VOCs) are the most frequently detected organic pollutants of groundwater in the United States. In fact, the VOCs are so ubiquitous that their analysis has been considered by the U.S. Environmental Protection Agency as a screening procedure to establish the need for more extensive characterization of groundwater samples from hazardous waste disposal sites. In the upstate region of New York (excluding Long Island), of approximately 570 groundwater contamination incidents reported by 1985, 98% involved either the volatile components of gasoline and petroleum or solvents and degreasers (NY State DEC, 1985).

Volatile organics may be transported in the subsurface as dissolved components in groundwater. However, by virtue of their volatility, VOCs will also exist in the gas phase of unsaturated porous media. As a result, volatile contaminants can be transported by advection and diffusion in the vapor phase. VOC transport processes are illustrated in Figure 1.

Experiment Description Students will

use soil gas sampling to prospect for a VOC that has leaked from a subsurface source into an unsaturated soil system. A rectangular “soil box” contaminated with a combination of liquid acetone, octane and toluene will be used. A soil with high organic

Spill, or Leaky Storage Tanks

Vadose Zone

VOC loss to the atmosphere

Groundwater Flow

VOC movement as a Non Aqueous Phase Liquid (NAPL)

VOC movement as a gas in soil pores

VOC movement as a solute dissolved in groundwater

Figure 1. Subsurface VOC transport processes. The vadose zone is the region of the soil profile in which some pores contain gas and are therefore, unsaturated.

120


content (potting soil) or low organic content (sand) may be used as the box filling material (porous medium). The VOCs will be identified and measured using a gas chromatograph (GC).


Calibration (Peak Times) Each compound will have a unique retention time in the gas chromatograph. The

time for each of the 3 VOC peaks can be obtained by injection of 100 µl head space samples from crimp cap sealed vials containing a small volume liquid acetone, octane, and toluene. Use the syringe technique described below. Analyze each compound 4 times (12 samples) using a gas chromatograph (see page 160 for information on using the gas chromatograph). These analyses will also serve to “calibrate” the GC by generating the peak area that corresponds to the saturated vapor concentration. Gas chromatogram peak areas may be assumed to be directly proportional to the mass of vapor injected.

Syringe technique for sampling vial headspace The purpose of this syringe technique is to minimize the effects of sorption to the

Teflon and glass surfaces in the syringe and to eliminate carryover of sample in the needle. Using separate needles to collect samples and to inject into the GC eliminates needle carryover of sample. 1) Remove GC needle. 2) Purge syringe 5 times with room air to remove any residual VOCs. 3) Put on sample needle. 4) Insert into sample bottle (with syringe at zero volume) 5) Fill syringe fully with gas, wait 4 seconds, and purge syringe contents back into

the source bottle (repeat 3 times). 6) Fill syringe and adjust to 100 µL. 7) Close syringe valve and remove syringe from sample vial and remove sample

needle. 8) Put on GC needle. 9) Instruct GC to measure sample (see page 160 for information on using GC

software). 10) Insert needle in injection port, open syringe valve, inject sample, hit start button

all as quickly as possible. 11) Remove syringe from the GC injection port.

Soil Moisture Content The dry weight of moist soil may be readily determined by placing ˜ 5 g moist soil

into a tarred aluminum weighing pan. Weigh the pan and its contents to obtain the soil’s wet weight, and place the pan into a 105oC oven for = 1 hour. Remove the soil

121


from the oven and place in a desiccator and allow it 5 minutes to cool. Weigh the cooled soil to obtain the dry weight. The moisture content is

wet weight - dry weight

Moisture Content =wet weight

10.1

Soil Density To determine the approximate density of the soil, ρsoil, place a weighed quantity of

dry soil (˜ 30 g) into a 100 mL graduated cylinder containing 60 mL water. Record the volume occupied by the water plus the soil.

2

soil = soil

total H O

MV V

ρ−

10.2

where ρsoil = ρb/θ, ρb= bulk density of the soil and θ = fraction of void volume.

Soil Gas Sampling See Table 1 for physical properties of the VOCs. See Tables 2, 3, and 4 (in the Lab

Prep Notes) for necessary reagents, equipment and GC method. Prior to the laboratory the instructor will create a “spill” of a VOC by injecting 10 mL of liquid of two or more NAPLs into the “soil box” to be sampled by students. During the lab students will analyze approximately 50 soil gas samples from the “soil box” using the syringe technique outlined below. Results from the soil box analyses may be mapped using units of concentration (g/m3).

Syringe technique for soil gas sampling 1) Remove GC needle. 2) Purge syringe 10 times with room air to remove any residual VOCs. 3) Put on sample needle. 4) Insert into soil bed (with syringe at zero volume). 5) Fill syringe and adjust to 100 µl. 6) Close syringe valve, remove syringe from soil bed and remove sample needle. 7) Put on GC needle. 8) Instruct GC to measure sample (using software).

Table 1. Physical data for octane, acetone, and

Octane Acetone Toluene Aqueous solubility

(g/m3)

0.6 very 515

Vapor Pressure (kPa)

1.88 (1.47) 24 3.8 (2.9)

H (kPa m3/mol)

300 0.0159 0.67

HGL (g/L)/(g/L)

123 0.0065 0.275

Molecular Formula

CH3(CH2)6CH3 CH3COCH3 C6H5CH3

Molecular weight

114.23 58.08 92.14

density (g/mL)

0.71 0.7857 0.8669

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9) Insert needle in injection port, open syringe valve, inject sample, hit the enter key (or OK) all as quickly as possible.

10) Remove syringe from the GC injection port.

Analysis of Soil Gas Sampling Students will use their analysis of VOC standards to obtain the corresponding GC

retention times and use this information to identify the unknown VOCs in the contaminated soil box. The vapor pressure and ideal gas law are used to estimate the mass of each compound present in the samples used to calibrate the GC.

PV

nRT

= 10.3

where n is the number of moles of the compound, P is the vapor pressure of the compound [kPa], V is the syringe volume [L], R is the Gas Constant (8.31 [ ] [ ]L kPa mol K⋅ ⋅ ), and T is the temperature of the gas in the syringe [K]. The relationship between peak area (as measured by the GC) and mass of the compound is determined from the calibration.

Soil gas concentrations should be reported and plotted as contour lines on a map of the soil box (see Figure 2 for an illustration).

Procedure (short version) 1) Instructor will demonstrate

syringe technique (be careful not to pull plunger out of barrel) and Gas Chromatograph technique.

2) Place ˜5 g of accurately weighed soil in oven to determine moisture content. (Weigh both the empty dish and the soil + dish.)

3) “Calibrate” GC by analyzing 4 samples for each VOC.

4) Take soil gas samples. 5) Convert the soil gas peak areas

to concentrations (g/m3). This data will be shared between groups. 6) Finish soil moisture content measurement (cool dry soil in desiccator and then

weigh). 7) Measure soil density using dry soil. 8) Pour waste potting soil into designated waste container. 9) Clean plasticware.

VOC NAPL source #2

VOC NAPL source #1

Sand

Sample grids

Figure 2. Students will prepare a map of the surface of their soil box. The map will show isoconcentration lines for each VOC.

123


Prelab Questions 1) How are the identities of the chromatogram peaks determined when using a gas

chromatograph? 2) Explain why different needles are used for sampling from source vials and

injecting the sample into the GC. Consider the temperature of the injection port (see Table 4) and the fact that these compounds absorb to most surfaces.

Data Analysis 1) Calculate the mass of each VOC in 100 µL of headspace. 2) Calculate the concentration of saturated vapor for each compound in g/m3. 3) Plot isoconcentration lines of the identified VOCs (expressed as gas concentration

in g/m3) on maps of the contaminated site (see figure 2 for example). Prepare a map for each compound showing isoconcentration lines. (The Excel 3-D surface plot with contour lines can be used. Note that the grid needs to have uniform distance between samples for the Excel 3-D surface plot to work correctly.)

4) Compare the saturated vapor concentration with the peak concentration observed in the “sand box.”

5) Calculate the soil moisture content and density.

References Ashworth, R. A., G. B. Howe, and T. N. Rogers. “Air-Water Partitioning Coefficients

of Organics in Dilute Aqueous Solutions.” J. Hazard. Mater. 18, p. 25-36, 1988. Council on Environmental Quality, "Contamination of Groundwater by Toxic

Organic Chemicals", 1981. Hwang, Y., J. D. Olson, and G. E. Keller, II, “Steam Stripping for Removal of

Organic Pollutants from Water. 2. Vapor-Liquid Equilibrium Data.” Ind. Eng. Chem. Res. 31, p. 1759-1768, 1992.

Mackay, D. and W. Y. Shiu, “A Critical Review of Henry’s Law Constants for Chemicals of Environmental Interest”, J. Phys. Chem. Ref. Data. 10, p. 1175-1199, 1981.

New York State Department of Environmental Conservation, "Draft Upstate New York Groundwater Management Program", N.Y.S.D.E.C., Division of Water, Draft Report WM P-94, January, 1985.

124


Lab Prep Notes

Setup

1) Prepare 1 soil box under fume hood.

2) Moisten the sand but not so much that there is standing water.

3) Pipette 10 mL of liquid acetone, octane, and toluene in sand box and record injection locations. This should be done in the morning before the lab exercise.

4) Dry approximately 100 g of potting soil for each group that will be used for density determinations.

5) Replace injection port septa on both GC’s.

6) Verify that GC’s are working properly by injecting gas samples from each VOC source bottle. If the baseline is above 30 (as read on the computer display) then heat the oven to 200°C to clean the column.

7) Verify that sufficient gas is in the gas cylinders (hydrogen, air, nitrogen).

8) Prepare VOC source vials that contain liquid acetone, octane, and toluene (they can be shared by two groups of students).

Class Plan

1) Setup uniform spreadsheets for data entry 2) Make sure spreadsheet is completely filled out by end of lab

Table 2. Reagents list

Description Source Catalog number

Octane Fisher Scientific 03008-1 Acetone Fisher Scientific O299-1 toluene Fisher Scientific T324-500

Potting soil Agway (remove large particles

by screening to 2 mm)



500 µl syringe w/ valve

Supelco 2-2272

side port needle Supelco 2-2289 1 mL syringe w/

valve Supelco 2-2273

Hp 5890 Series II GC

Hewlett-Packard 5890A

Sep purge-packed/FID

Hewlett-Packard option 600

1/8" column adapter

Hewlett-Packard option 095

pressure regulators Hewlett-Packard L43 RS232C board Hewlett-Packard option 560

Nitrogen, Air, and Hydrogen gas

General Stores

Wrist action Shaker Fisher Scientific 14-260 Desiccator Fisher Scientific 08-642-15

Vials Supelco 3-3111 Aluminum crimp

tops Supelco 3-3220

Septa Supelco 3-3200 Crimping tool Supelco 3-3280

125


Table 4. Gas chromatograph conditions

gas pressure flow carrier (N2) 320 kPa 15 mL/min

Air 230 kPa 300 mL/min Hydrogen 130 kPa 45 mL/min

temperatures °C

oven (isothermal) 100 Injector 250

FID 250

Column Supplier Catalog number

Supelcowax 10 30 meters

Supelco 2-5301

Run length of 66 seconds with octane, acetone, and toluene at 0.57, 0.63, 0.96 minutes respectively. Maximum sample volume is about 100 µl. Larger samples can lead to a significant broadening of the peak.

Syringe clean up Disassemble and heat syringes to 45°C overnight to remove residual VOCs. Place

syringe barrels upside down on top of openings above fan in oven to facilitate mass transfer.

126


Volatile Organic Carbon Sorption to Soil

Introduction Volatile organic carbon compounds (VOCs) can exist as vapors, non-aqueous

phase liquids, dissolved in water, or sorbed to surfaces. Sorption is the term used to refer to the binding reactions between organic pollutants and the subsurface medium. Sorption slows the rate of transport of both dissolved and volatile organic pollutants. Laboratory experiments will be performed to evaluate the sorption of acetone, hexane, and octane vapors to a soil and to estimate the extent to which VOC transport is slowed by binding to the soil media.

Theory

Gas-Liquid Partitioning The equilibrium between gas and solution is described by the following reaction:

Liquid GasC C↔ 11.1

The equilibrium constant ( GLH ) for the reaction is given by:

G GL

L

CH

C= 11.2

where CG is the concentration in the gas phase (Example units: g/m3) and CL is the concentration in the aqueous phase (g/m3)

Alternately, the liquid-vapor equilibrium is sometimes expressed as:

G

L

PH

C= 11.3

where PG is the partial pressure of the gas (atm), and

GL

HH

RT= 11.4

where R is the universal gas constant 3

58.21 10m atm

xmol K

− ⋅ ⋅

and T is the absolute

temperature (oK). Both H and G

LH are referred to as “Henry's law constants” and may be viewed conceptually as distribution coefficients for gases between the aqueous solution phase and the vapor phase. All other factors being equal, VOCs with higher Henry's law constants will have a greater fraction of their total mass in the gas phase of an unsaturated porous medium.

127


Liquid-Solid Partitioning The sorption of organic pollutants that are dissolved in water onto soils and aquifer

materials also may be described in many cases by a distribution coefficient ( SLK ):

SL

L

KCΓ

= 11.5

where Γ is the mass of solute sorbed per mass of solid. Equation 11.5 predicts that the amount of pollutant bound to the soil (Γ) will

increase linearly with the concentration in the aqueous phase (CL). Any relationship between the amount bound to the soil and the concentration in solution applies at a constant temperature and is referred to as an “isotherm.” Equation 11.5 is an example of a “linear isotherm.” The success of the linear isotherm in describing sorption of nonionic organic pollutants in saturated soils has been remarkable. Linear isotherms have been found to describe sorption of a wide array of nonionic compounds onto sediments and soils (Karickhoff et al. 1979, Chiou et al. 1979).

Many investigations have demonstrated that the distribution coefficient ( SLK ) for

sorption of a single organic contaminant onto a variety of soil materials can be related to the organic content of the sorbent. This observation permits the definition of an organic normalized partitioning coefficient (Koc):

? f

SL

ococ

KK = 11.6

where foc is the weight fraction of organic carbon in the soil. Koc values for a range of different organic compounds have been shown to be

related to their octanol-water partitioning coefficients (Kow) and also to their aqueous solubilities (Karickhoff, 1984). An important implication of these results is that sorptive distribution coefficients of organic pollutants in water saturated aquifers ( S

LK ) may be predicted given knowledge of the organic content of the soil (foc) and the octanol-water partitioning coefficient (Kow) of the contaminant or a related parameter such as its aqueous solubility.

Gas-Solid Partitioning Sorption of gases is frequently described using the classic equation developed by

Brunauer, Emmett and Teller (1938), i.e., the BET equation:

0 0

0 0

M

B

P P P PB

P P

Γ=

Γ − −+

11.7

where Γ is the amount of sorbed gas per unit of surface (with units such as moles/m2 or µg/g if the surface area is not known), ΓM is the amount of sorbed gas corresponding to monolayer surface coverage, P is the partial pressure of the sorbed

128


gas, Po is its saturated vapor pressure, and B is a parameter related to solute binding intensity; more specifically:

v

kTB eε ε−

= 11.8

where ε is the energy of the adsorbate vapor-adsorbent surface interaction (cal/mole), εv is the vaporization energy of the organic (cal/mole), k is the Boltzmann constant, and T is the absolute temperature (oK).

To the extent that it is valid in soil, the BET equation predicts vapor sorption isotherms to be nonlinear. Nonlinear isotherms have been observed for sorption of organic vapors onto dry soils at high vapor concentrations by several investigators (Chiou, 1990; Rhue, et al., 1988; and Ong and Lion, 1991c). Under conditions of low vapor pressure, P << Po, the BET equation reduces to:

0M

BP

BP

Γ=

Γ +

11.9

which is the “Langmuir adsorption equation” that applies to monolayer limited adsorption.

The BET equation further reduces to a linear isotherm when B << (Po/P).

0M

PB

P Γ

= Γ 11.10

Figure 1 illustrates linear, Langmuir and BET isotherms that share a common set of B and Γm values. Linear sorption isotherms for vapors are a reasonable expectation at low vapor concentrations (Ong and Lion, 1991a).

At higher vapor concentrations, surface site limitations and the phenomenon of vapor condensation at the surface (for which the BET model attempts to account) will result in nonlinear VOC sorption isotherms. Results obtained at Cornell (Ong

0.30.20.10.00

4000

8000

12000

16000

20000

Γ

P/Po

LinearBET

Langmuir

B = 1,000max = 10,000Γ

A

1.00.80.60.40.20.00

100000

200000

300000

Γ

P/Po

BET

Langmuir

B

Figure 1. Linear, Langmuir and BET sorption isotherms for the case where Γm = 10,000 and B = 1,000. A) Isotherms for P/Po < 0.3. B) Isotherms for P/Po up to 1.0.

129


et al., 1991; Ong and Lion, 1991c) have confirmed that condensation of organic vapors will occur at high vapor pressures in moist porous media. Condensed water and organic vapors compete for the available pore space. Since soils are generally wet prior to the introduction of organic contaminants, vapor condensation is expected to be limited to the pore volume not occupied by water. The extent to which organic vapor condensation is significant will, therefore, be a function of the soil moisture content, and the relative vapor concentration (P/P0).

Since the unsaturated zone in an aquifer will typically contain condensed water, description of the sorption of organic vapors in the unsaturated zone must, at a minimum, consider a binary mixture of the organic and water vapor. The organic vapor of concern may accumulate in the unsaturated zone through at least three processes: (a) by direct sorption from the vapor phase, including vapor sorption to dry mineral surfaces (if present), vapor sorption at the gas-water interface, and vapor condensation, (b) by solubilization in condensed pore water as governed by Henry's Law (equation 11.2 or 11.3), and (c) by sorption from condensed pore-water solution onto the soil (as governed by equation 11.5). Since, at very low vapor pressures, a linear isotherm is expected to govern vapor sorption, we may write a linear isotherm in terms of the gas concentration:

SG GK CΓ = ? 11.11

where the magnitude of SGK depends on the sorbent’s moisture content.

The relative contributions to Γ of processes such as vapor dissolution into soil moisture and sorption at the liquid-air interface can be assessed through the use of a mass balance. The total mass of vapor sorbed onto the soil, under equilibrium conditions, can be viewed as being distributed between: (Mass sorbed at the solid-liquid interface) + (Mass dissolved in the liquid phase) + (Mass sorbed at liquid-air interface + condensation)

= = + +s L LM X C V ωΓ 11.12

where Ms is the mass of the sorbent (soil or other porous media) and ω is a lumped parameter that includes the effects of water surface sorption and condensation.

From equation 11.5 for liquid-phase sorption: SL S LX K M C= where Γ = X/Ms and

Ms is the mass of the sorbent. Also, from Henry's Law: gG L LC H C= . Substituting

these two relationships and equation 11.11 into equation 11.12 gives:

S

S L LG G G

G SL S L

K VK

C MH M Hϖ

= + + 11.13

If the density of water on the soil surface is expressed as 2 2

/H O H O LM Vρ = , the

term VL/Ms becomes 2

2

H O

s H O

M

M ρ

. Assuming water surface sorption and condensation

are negligible (ω ˜ 0), then for high moisture content equation 11.13 will plot as a

130


straight line, with the ordinate intercept equal to the contribution of aqueous phase sorption to vapor-phase partitioning ( /S G

L LK H ).

Moisture Content %

100

SS LG G G

L L

KK

H H= + 11.14

Equation 11.14 indicates that the linear vapor distribution coefficient, SGK , can be

predicted from the saturated distribution coefficient and the Henry’s law constant for a given VOC. Experiments by Ong and Lion (1991a) have shown that such predictions are reasonable as long as the moisture content of the soil is high enough to ensure that the VOC that is dissolved in sorbent-bound water behaves as if the liquid were comparable to the water in a bulk aqueous phase. In general, a moisture content equivalent to an average surface coverage of ˜ 5 layers of water molecules is adequate for this assumption to be obeyed (Ong and Lion, 1991a). Many soil ambient moisture contents are in excess of this limiting value.

Pollutant Transport in Porous Media The advective dispersion equation is used to describe pollutant movement in

porous media. For one-dimensional (ex., horizontal) transport of a conservative ( 0S

LK = ) pollutant:

2

2 -U E

C C Ct x x

∂ ∂ ∂= +

∂ ∂ ∂ 11.15

where t is time, x is distance, U is the groundwater pore velocity and E is the macroscopic dispersion coefficient (Freeze and Cherry, 1979).

Since volatile organic pollutants react with the surfaces of the porous media through which the contaminant flows, equation 11.15 must be modified to account for

the sorption reaction by addition of the term: b

tρθ

− ∂Γ∂

where bρ is the bulk density of

the porous medium (g/cm3), and θ is the volumetric moisture content (volume of liquid per unit bulk volume of the porous medium). θ is equal to the porosity, φ, in a saturated porous medium.

From the chain rule, b b Ct C t

ρ ρθ θ

∂Γ ∂Γ ∂=

∂ ∂ ∂and for a linear isotherm, S

LKC

∂Γ=

∂.

Therefore, the advection-dispersion equation for a compound that experiences sorptive binding to the soil matrix becomes:

= - + ∂∂

∂∂

∂∂

- ∂∂

Ct

uCx

EC

xK C

tb L

S2

2

rq 11.16

1+ = - +∂∂

FHG

IKJ

∂∂

∂∂

Ct

Ku

Cx

EC

xb L

Srq

2

2 11.17

The retardation factor, R, for a pollutant in soil is defined as:

131


1+S

b LKR

ρθ

=

11.18

Therefore the advective dispersion equation modified for sorptive binding becomes:

2

2

- = +

C u C E Ct R x R x

∂ ∂ ∂∂ ∂ ∂

11.19

From equation 11.19 it is apparent that the velocity and dispersion of a sorbed compound will be reduced by the magnitude of retardation factor, R. So,

pore water velocity

= 1contaminant velocity

R ≥ 11.20

Note that R may be determined directly from knowledge of the medium properties ( bρ and θ) and from the distribution coefficient for sorption ( S

LK for water saturated

media or SGK for a vapor).

Interestingly, equation 11.19 may also be applied to describe vapor movement in a gas chromatograph (GC). GC columns are selected to ensure that the components of a vapor mixture will be separated (by virtue of their different retardation factors) by the time they arrive at the GC detector (situated at the end of the column).

In the absence of pressure gradients, transport of vapors will occur primarily through the process of diffusion and equation 11.19 reduces to:

2

2s

bulk

EC Ct E x

∂ ∂=

∂ ∂ 11.21

where Es is the effective diffusion coefficient of the VOC in the porous media. Vapor diffusion coefficients in unsaturated porous media are different from those

in a bulk gas phase because the vapor must follow a tortuous path to move through the open pores. The relationship proposed by Millington and Quirk (1961) is commonly used to correct vapor diffusion coefficients for the conditions in the soil media.

bulks Ea

E 2

3/10

φ= 11.22

where Ebulk is the vapor diffusion coefficient in air, a is the volumetric air content of the porous medium (volume of gas per unit bulk volume of medium), and φ is the porosity (a + θ = φ).

Analysis of the Unsaturated Distribution Coefficient ( SGK )

A mass balance calculation will be used to determine the unsaturated vapor distribution coefficient ( S

GK ). After equilibration the VOC mass will be distributed between the vapor phase and the solid phase (sorbed VOC).

132


= +VOC G G sM C V MΓ 11.23

where sMΓ is the mass of sorbed VOC and equals SG S GK M C (from equation 11.11).

In control bottles there is no sorbent so the VOC mass must reside entirely in the vapor phase:

= UC UC UCVOC G GM C V 11.24

where the subscript uc indicates that volumes and concentrations are those measured in the unsaturated control bottles.

Setting equations 11.23 and 11.24 equal to one another (the mass of VOC was the same for all vials) and rearranging gives:

= UC

SVOC G G G S GM C V K M C+ 11.25

Solving for SGK

UCVOC G GSG

S G

M C VK

M C

−= 11.26

Using the measured soil moisture contents and values of SGK , students may check

the validity of equation 11.14.

Unsaturated Mass Fraction Distribution The total mass of the VOC is distributed between the gas and sorbed phases

(equation 11.23).

1=ƒ +ƒG S 11.27

where ƒG is the fraction of the VOC mass in the gas phase and ƒS is the fraction of the VOC mass sorbed to the soil. The relationship between the fraction of VOC in each phase is obtained from the definition of the unsaturated distribution coefficient (equation 11.11).

ƒ

= ƒ

SS G S

G G

K MV

11.28

Thus we have two equations in two unknowns. Solving we obtain

1

ƒ =1

G SG S

G

K MV

+ 11.29

1

ƒ =1

SG

SG S

VK M

+ 11.30

where

133


= SG vial

soil

MV V

ρ− 11.31

Determination of SGK requires a measurement of the fraction sorbed given

measurements of the total mass added and the mass in the gas phase. This analysis becomes inaccurate as the magnitude of ƒS decreases and approaches the coefficient of variation of

UCVOCM .

Analysis of the Saturated Distribution Coefficient ( SLK )

A mass balance calculation will be used to determine the saturated vapor distribution coefficient ( S

LK ). The calculation assumes that students can reproduce the introduction of the mass of VOC (MVOC) into each sample bottle. After equilibration the VOC mass can be distributed between the vapor phase the aqueous phase and the solid phase (sorbed VOC).

= + + VOC G G L L SM C V C V MΓ 11.32

where ΓMs is the mass of sorbed VOC and equals SL L SK C M (from equation 11.5).

In saturated control bottles there is no sorbent so the VOC mass must just be distributed between the vapor phase and the aqueous phase:

= + SC SC SC SC SCVOC G G L LM C V C V 11.33

where the subscript sc indicates that volumes and concentrations are those measured in the control bottles. Henry’s law (equation 11.2) can be used to eliminate

scLC .

C = SC

SC S SC

LVOC G G G

L

VM C V

H

+ 11.34

Equation 11.34 can be used to obtain an estimate of Henry’s law constant by assuming that the mass of VOC added is the same for the vials with and without water. Solving for G

LH and substituting UCVOCM for

SCVOCM we obtain:

= SC SC

UC SC SC

L GGL

VOC G G

V CH

M V C− 11.35

Equation 11.32 can now be used to obtain an estimate of SLK . The mass of VOC

added to the bottles was the same for all vials. Thus ( VOCM ) in equation 11.32 is equal to

SCVOCM . In addition, CL and CG are interrelated through Henry’s law (equation 11.2). Substituting into equation 11.32 gives:

+ = +

sc

LG GVOC G G L S SG G

L L

C CM C V V M K

H H 11.36

Solving for SLK

134


sc

GS GLL VOC G G LG

S G L

CHK M C V V

M C H

= − −

11.37

Measured VOC headspace concentrations are substituted into equation 11.37. Values for Henry’s law constants are given in Table 1.

SCVOCM is obtained from equation 11.34. In vials containing soil, VG is determined by subtracting both VL (50 mL) and the volume of the sorbent from the total vial volume. The volume of the sorbent may be calculated from the sorbent weight and density as:

ssoil

soil

MV

ρ= 11.38

It is instructive to calculate the phase distribution of each VOC in bottles that contain no soil. The fraction (ƒ) of VOC mass in the gas phase is given by

= =G G G

LG G L LGGL

C V Vf

VC V C V VH

+ + 11.39

Assuming the total volume of the bottle is 120 mL and 50 mL of liquid are added, then

70

=5070 G

L

f

H+

11.40

ƒ values for octane, acetone, and toluene are therefore 0.994, 0.009, and 0.278 respectively indicating that only toluene has a significant mass fraction in both the gas and aqueous phases. In the absence of strong sorption by soil, octane will reside primarily in the gas phase and acetone will reside primarily in the aqueous phase. Determinations of S

LK for these two compounds may therefore not be feasible using the headspace technique.

Saturated Mass Fraction Distribution The total mass of the VOC is distributed between the gas, sorbed, and liquid phases

(equation 11.32).

1= + +G S Lf f f 11.41

where Lf is the fraction of the VOC mass in the liquid phase. The relationship between the fractions of VOC in the solid and liquid phases is obtained from the definition of the saturated distribution coefficient (equation 11.5).

= S

S L S

L L

f K Mf V

11.42

The relationship between the fractions of VOC in the gas and liquid phases is obtained from the definition of the Henry’s law constant (equation 11.2).

135


= G

G L G

L L

f H Vf V

11.43

Thus we have three equations in three unknowns. Solving we obtain

=+

LL S G

L L S L G

Vf

V K M H V+ 11.44

=+ +

SL S

S S GL L S L G

K Mf

V K M H V 11.45

=+ +

GL G

G S GL L S L G

H Vf

V K M H V 11.46

where

= SG vial L

S

MV V V

ρ− − 11.47

Determination of SLK requires a measurement of the fraction sorbed given

measurements of the total mass added, the mass in the gas phase, and knowledge of the Henry’s law constant. This analysis becomes inaccurate when ƒS decreases and approaches the coefficient of variation of

SCVOCM .

Experimental procedures VOC distribution coefficients will be determined using crimp top vials sealed with

Teflon backed septa and aluminum crimp caps. VOCs will be analyzed by students using a gas chromatograph (GC).

A data table for preparation of the necessary vials is shown in Table 4. The data table is also available in spreadsheet form as “isotherm calculator.”

Analysis of the Unsaturated Distribution Coefficient ( SGK )

The distribution coefficient for three VOCs (acetone, octane, and toluene) with the unsaturated soil medium ( S

GK ) will be determined using the headspace method of Peterson et al. (1988). Students will prepare 120 mL vials (actually 121.5 ± 0.5 mL) in triplicate containing 0 and 20 g of moist soil (6 vials total). The weight of replicates need not be precisely matched, but the weight of soil should be measured accurately. Seal the vials using Teflon-backed septa (the shiny Teflon side faces the vial contents) and aluminum crimp cap.

The 3 VOCs are added to the vials by introduction of 1 mL of the saturated vapor taken from the headspace of “source vials” (crimp capped vials containing liquid octane, toluene, and acetone). Use a dedicated syringe for each VOC and leave dedicated needles in the source bottles.

The vials should be placed on a wrist action shaker to continue agitation for = 30 minutes to permit the VOC time to sorb to the soil medium. After equilibration, vials

136


are removed from the wrist action shaker and the head space sampled using a 500 µL gas tight syringe. Use the syringe technique as described previously for sampling vial headspace.

Analysis of the Saturated Distribution Coefficient ( SLK )

The saturated distribution coefficient ( SLK ) for three VOCs (acetone, octane, and

toluene) with the soil medium will be determined using the headspace method of Garbarini and Lion (1985) as modified by Peterson et al. (1988). These vapors are selected to demonstrate the effect of VOC Henry's law constant on VOC phase distribution (see Table 1).

Students will prepare 120 mL vials (actually 121.5 ± 0.5 mL) in triplicate containing 0 and 20 g of potting soil (6 vials total). The weight of replicates need not be precisely matched, but the weight of soil should be measured accurately. Soil density should be separately measured as described below. If moist soil is used students should also determine the dry weight unless the instructor provides this information (see below). Into each vial students will add 50 mL of tap water.

Seal the vials using a Teflon-backed septa (the shiny Teflon side faces the vial contents) and aluminum crimp cap. Octane and toluene are then added to the vials by introduction of 1 mL of the saturated vapor taken from the headspace of “source vials” (crimp capped vials containing liquid octane and toluene). Use different needles for collecting and delivering the VOC to reduce the transfer of VOC on the needles. The needle used to collect the VOC from the source bottle can be left in the source bottle and simply attached to the syringe when needed. Acetone is added by introduction of 200 µl of the liquid compound using a gas tight syringe.

The vials should be placed on a wrist action shaker to continue agitation for = 30 minutes to permit the VOC time to sorb to the soil medium. After equilibration, vials are removed from the wrist action shaker and the head space sampled using a 500 µL gas tight syringe. To sample the vial headspace, use a 100 µL sample and the syringe technique described previously.

Procedure (short version) 1) Weigh soil and add to isotherm vials (6 vials with 20 g). 2) Add 50 mL tap water to 6 vials (3 each with 0 and 20 g soil). 3) Seal 12 vials. 4) Add octane, toluene, and acetone to all vials (1 mL gas from source vials for all

VOC’s except 200 µL liquid acetone for vials with tap water). 5) Place vials on shaker for 30 minutes.

6) “Calibrate” GC by analyzing 4 100-µL samples for each VOC. 7) Take vials off of shaker. 8) Measure VOC concentrations for each vial and record peak areas in spreadsheet. 9) Reanalyze the VOC concentrations for any vials for which anomalous data was

obtained. 10) Remove vial caps.

137


11) Pour waste potting soil into designated container. 12) Wash vials.

Prelab Questions 1) Why does the determination of S

GK become inaccurate as the magnitude of ƒS decreases and approaches the coefficient of variation of

UCVOCM ?

2) What conditions are necessary to obtain linear isotherms for gas/solid partitioning of organic vapors?

3) When can equation 11.14 be used to predict the relationship between liquid/solid partitioning and gas/solid partitioning based on soil moisture content and Henry’s law constant?

Data Analysis

A Note on Units Express mass of VOC in grams (as measured by the GC). Express concentrations

in g/mL. Remember to account for the fact that the syringe volume for GC analysis is 100 µL Express all volumes in mL. 1) Estimate the mass of each VOC added to the unsaturated sample vials based on

UCVOCM (from equation 11.24). Report mean and coefficient of variation (standard deviation/mean) for each VOC.

2) Calculate the unsaturated vapor distribution coefficient ( SGK ) using equation

11.26. Report a single mean and coefficient of variation for each VOC. 3) Calculate the mass fraction associated with the soil and gas phases under

unsaturated conditions for each of the VOCs. Use equations 11.29, 11.30, and 11.31. Assume Ms = 20 g and bottle volume is 121.5 mL. Use the average S

GK calculated in 2 above. Compare the coefficient of variation of

UCVOCM to the mass

fractions to evaluate which SGK determinations are potentially accurate. Create a

stacked bar graph of the mass fractions for each VOC.

4) Estimate the Henry’s law constant ( GLH ) for octane and toluene using equation

11.35 (different masses of acetone were used for the saturated and unsaturated vials and thus equation 11.35 can not be used for acetone). Report mean and coefficient of variation. Compare your results to the Henry’s law constants reported in the table on page 121.

5) Estimate the mass of each VOC added to the saturated sample vials based on

SCVOCM (from equation 11.34) using tabulated Henry’s constants (see table on page 121). Report mean and coefficient of variation for each VOC.

6) Calculate the saturated vapor distribution coefficient ( SLK ) using equation 11.37.

Report a single mean and coefficient of variation for each VOC. Use tabulated Henry’s constants reported in the table on page 121.

138


7) Calculate the mass fraction associated with the soil, gas, and water phases under saturated conditions using equations 11.44 - 11.47. Assume Ms = 20 g, VL = 50 mL, and bottle volume is 121.5 mL. Use the average calculated S

LK and the Henry’s law constants reported in the table on page 121. Compare the coefficient of variation of

SCVOCM to the mass fractions to evaluate which SLK determinations

are potentially accurate. Create a stacked bar graph of the mass fractions for each VOC.

8) Calculate SGK for toluene using equation 11.14 and compare with the measured

value of SGK .

9) Assuming a typical soil porosity (θ) of 0.33 and bulk density (ρb) of 1.7 g/cm3, equation 11.18 may be used to calculate the retardation of dissolved VOCs and VOC vapors. Assuming a pore water velocity of 1 m/day, how long will it take for the dissolved toluene to be transported a distance of 100 m? Include a range of the estimate based on 1 standard deviation.

10) If toluene is removed by withdrawing vapor at a velocity of 100 m/day, how long will it take toluene vapor to travel 100 m? [Note in this case S

GK replaces SLK and

a replaces θ in equation 11.18. Assume the soil voids are filled with gas so a = θ = 0.33.] Include a range of the estimate based on 1 standard deviation.

11) Discuss how the results of this experiment would guide you in remediating a site contaminated with toluene, acetone, and, octane.

References Brunauer, S., P.H. Emmett and E. Teller, "Adsorption of Gases in Multimolecular

Layers", J. Amer. Chem. Soc., 60, p. 309, 1938. Chiou, C.T., L.J. Peters and J.H. Freed, "A Physical Concept of Soil-Water Equilibria

For Nonionic Organic Compounds", Science, 206, p. 831, 1979. Chiou, C.T., "Roles of Organic Matter, Minerals and Moisture in Sorption of

Nonionic Compounds and Pesticides by Soils", in Humic Substances in Soil and Crop Sciences; Selected Readings, P. MacCarthy, C.E. Clapp, R.L. Malcolm and R.R. Bloom (Eds.), Am. Soc. of Agron. & Soil Sci. Soc. of Am, Madison, WI, pp. 111-160, 1990.

Freeze, R.A. and J.A. Cherry; Groundwater, Prentice Hall, Inc.; Englewood Cliffs, NJ, 604 pp., 1979.

Karickhoff, S.W., D.S. Brown and T.A. Scott, "Sorption of Hydrophobic Pollutants on Natural Sediments", Water Res., 13, p. 241-248, 1979.

Karickhoff, S.W., "Organic Pollutant Sorption in Aquatic Systems", J. Hydraulic Engrg., 110(6), p. 707, 1984.

Millington, R.J., J.M. Quirk, “Permeability of porous solids”, Trans. Faraday Soc. 57, p. 1200-1207, 1961.

Ong, S.K., and L.W. Lion, "Sorption Equilibria and Mechanisms for Trichloroethylene onto Soil Minerals", J. Env. Qual., 20(1), p. 180-188, 1991a.

139


Ong, S.K., and L.W. Lion, "Effects of Soil Properties and Moisture on the Sorption of TCE Vapor", Water Resources Research, 25(1), p. 29-36, 1991b.

Ong, S.K., and L.W. Lion, "Trichloroethylene Vapor Sorption onto Soil Minerals at High Relative Vapor Pressure", Soil Sci. Soc. Am. J., 55(6), p. 1559-1568, 1991c.

Ong, S.K., S.R. Lindner and L.W. Lion, "Applicability of Linear Partitioning Relationships for Organic Vapors onto Soil Minerals", in: Organic Substances and Sediments in Water, R.A. Baker (Ed.), Lewis Publ., Chelsea, MI, pp. 275-289, 1991.

Rhue, R.D., P.S.C. Rao and R.E. Smith, "Vapor Phase Adsorption of Alkylbenzenes and Water on Soils and Clays", Chemosphere, 17, p. 727-741, 1988.

Additional References Relevant to Data Reduction Garbarini, D.R. and L.W. Lion, “Evaluation of sorptive partitioning of nonionic

organic pollutants in closed systems by headspace analysis,” Env. Sci. & Tech., 19(11), 1122-1128 (1985).

Gossett, J.M. “Measurement of Henry's law constants for C1 and C2 chlorinated hydrocarbons,” Env. Sci. & Tech., 21(2), 202 (1987).

Peterson, M.S., L.W. Lion, and C.A. Shoemaker, “Influence of vapor phase sorption and diffusion on the fate of trichloroethylene in an unsaturated aquifer system.” Env. Sci. & Tech. 22(5), 571-578, (1988).

140


Symbol List Symbol Definition

GLH Henry’s Constant

SLK Liquid/solid partitioning coefficient

SGK Gas/solid partitioning coefficient

B Solute binding intensity C Concentration E Dispersion coefficient f Mass fraction k Boltzmann constant

Koc Liquid/organic carbon partitioning coefficient

Kow octanol-water partitioning coefficients M Mass P Partial pressure P0 Saturated vapor pressure R Universal gas constant R Retardation factor T Absolute Temperature(oK). T Temperature t Time U Groundwater pore velocity V Volume X Mass sorbed at the solid-liquid interface x Distance Γ Mass of solute sorbed per mass of solid ε vapor-adsorbent surface interaction εv vaporization energy of the organic

φ Porosity θ Volumetric moisture content ρ Density of water ω Mass sorbed at liquid-air interface +

condensation

Subscript Definition b Bulk G Gas phase L Aqueous phase M Monolayer oc Organic carbon S Soil phase or sorbent sc Saturated control uc Unsaturated control

VOC Volatile organic carbon

141


Lab Prep Notes For equipment list and gas

chromatograph method see page 124.

Setup

1) Replace injection port septa on all GC’s.

2) Verify that GC’s are working properly by injecting gas samples from each VOC source bottle. If the baseline is above 30 (as read on the computer display) then heat the oven to 200°C to clean the column.

3) Verify that sufficient gas is in the gas cylinders (hydrogen, air, nitrogen). 4) Verify that sufficient vials/aluminum crimp tops and Teflon seals are available. 5) Prepare VOC source vials that contain liquid acetone, octane, and toluene (they

can be shared by two groups of students). 6) Fill repipet dispensors with tap water and place on each lab bench.

Table 6. Each group of students requires the following syringes

Octane (gas) 1 mL Toluene (gas) 1 mL Acetone (gas) 1 mL

Acetone (liquid) 0.5 mL Source vial (gas) 0.5 mL Isotherm (gas) 0.5 mL

Syringe clean up Disassemble and heat syringes to 45°C overnight to remove residual VOCs. Place

syringe barrels upside down on top of openings above fan in oven to facilitate mass transfer.

Table 5. Reagents list

Description Source Catalog number

Octane Fisher Scientific 03008-1 Acetone Fisher Scientific O299-1 toluene Fisher Scientific T324-500

Potting soil Agway (remove large particles

by screening to 2 mm)

142


Enhanced Filtration

Introduction Slow sand filters have been used to remove particles from drinking water since the

early 1800's. Although slow sand filtration is an old technology, the mechanisms responsible for particle removal are not well understood. Because slow sand filter performance gradually increases with time, it has often been assumed that the growth of biofilms is responsible for the gradual improvement in filter performance. Research conducted at Cornell suggests that biofilms are not responsible for significant particle removal and that most particles are removed by physical-chemical mechanisms.

The particles that are captured on slow sand filters have been shown to significantly improve filter performance (Weber-Shirk and Dick, 1997). More recent research has shown that a filter aid can be extracted under acid conditions from particles harvested from Cayuga Lake or from Cayuga Lake sediment. The filter aid has been shown to greatly enhance bacteria removal. The filter aid is soluble at very low pH, and forms floc at neutral pH. This naturally occurring filter aid may be able to improve rapid sand filter performance.

Theory In new slow sand filters with clean filter media, particles are initially removed by

attaching to the filter media. However, as the filter media begins to be covered with removed particles, particles begin to attach to previously removed particles. If particle-particle interaction is more favorable than particle-media interaction then particle removal efficiency increases as the media becomes covered with particles. This improvement in filter performance with time is commonly observed in slow sand filters and is referred to as filter “ripening.” Filter ripening often takes several weeks to several months for new slow sand filters. Slow sand filters that operate with pristine water sources may never achieve efficient particle removal because the lack of particles in the source water results in a sparse coating of the filter media.

Potential mechanisms of particle removal by slow sand filters are summarized in Figure 1. Physical-chemical removal mechanisms are responsible for most of the particle removal that occurs in slow sand filters. The one exception is that suspension-feeding nanoflagellates attached to the filter media can capture a significant fraction of bacteria (Weber-Shirk and Dick, 1999). Thus, bacteria removal by suspension feeding predators is significant provided the influent bacteria concentration is sufficient to maintain a large predator population. Biofilms on the filter media have not been shown to significantly increase particle removal.

Straining of bacteria-sized particles by the filter media and attachment of bacteria-sized particles to the filter media were shown to not be significant because the removal of bacteria by a clean filter column was negligible (Weber-Shirk and Dick, 1997). It is possible that straining becomes significant as filters clog and pores become smaller. Attachment of particles to previously removed particles is considered likely.

143

Enhanced Filtration

Physical-chemical filter ripening may be the result of the changes in pore geometry that enhance straining or the modification of filter media surfaces that enhance the ability of particles to attach. Decreasing the pore size to enhance straining is not a reasonable way to improve particle removal because the head loss through the filter increases rapidly as the pore size decreases. Thus, the best way to enhance physical-chemical ripening is to modify filter media surfaces. The filter aid may act by coating the surface and providing more favorable attachment sites.

Filtration theory suggests that particle removal will be first order with respect to depth if the filter media is homogeneous (Iwasaki, 1937). In equation form the relationship between particle concentration, C, and depth is given by

dC

Cdz

λ= − 12.1

where λ is the filter coefficient with units of [1/L]. Setting appropriate integration limits

0 0C

C L

dCdz

Cλ= −∫ ∫ 12.2

where L is the depth of the filter bed and Co is the influent particle concentration and integrating gives:

0

lnC

LC

λ= − 12.3

Suspensionfeeders

Grazers

Attachment tobiofilms

Capture bypredators

to medium

to previouslyremovedparticles

by medium

bypreviouslyremovedparticles

Straining(fluid and

gravitationalforces)

Attachment(electrochemical

forces)

Physical-Chemical

Biological

ParticleRemoval

Mechanisms

Figure 1. Particle removal mechanisms that potentially could be operative in slow sand filters.

144


Equation 12.3 can be used to evaluate the filter coefficient, λ. A list of previously measured filtration constants is given in Table 1.

Filtration theory suggests that filter performance would be optimal if the filter aid were applied uniformly throughout the filter. Uniform application is difficult, however, because the filter aid will be captured first order with respect to depth if the filter aid is applied using normal down flow operation. It may be possible to apply the filter aid during a gentle backwash thus enabling the filter aid to distribute more uniformly. Application techniques that optimize the filter aid distribution require further study.

Previous Research Results Previous research (Weber-Shirk and Dick, 1997) has shown that Cayuga Lake

water particles can enhance filter performance and thus Cayuga Lake particles (CLP) from the Bolton Point Water Treatment Plant sedimentation basin were tested. Three filters were treated with 30 mL of concentrated CLP suspension from the Bolton Point Water Treatment Plant. One filter had the CLP mixed throughout the filter bed, one filter had the CLP mixed throughout the top 2 cm of the filter bed, and one filter had the CLP applied only to the top of the filter bed. The three application techniques were used because particles may improve filtration efficiency by providing surfaces to which bacteria attach more readily or because the pores within the sediment are smaller and thus more effective at straining particles. The filter with the particles distributed throughout the filter bed performed the best with approximately 99% removal of kaolin compared with 96% removal for the filter with the CLP on top of the filter bed. This result suggested that kaolin was being removed by attaching to CLP rather than by straining. CLP from the Bolton Point facility contain alum and possibly other polymers used in the water treatment process.

Previous research also indicated that an acid treatment of Cayuga Lake sediment dissolves species that flocculate and attach to filter media at neutral pH. This Cayuga Lake Sediment Extract (CLSE) has been shown to rapidly ripen slow sand filters and achieve up to 6 log (99.9999%) removal of E. coli. The CLSE has also been shown to enhance E. coli removal at rapid sand filtration rates.

Table 1. Typical values of filter coefficients adapted from (Tien and Payatakes, 1979).

Filter medium

Grain size (mm)

particle type

particle size (µm)

approach velocity (cm/hr)

λ (1/cm)

Calcium carbonate

?? Ferric floc

10 500 0.1

Calcium carbonate

?? Ferric floc

10 1000 0.044

Anthracite 0.77 Quartz powder

2-22 500 0.064

Sand 0.54 Chlorella

5 500 0.34

Sand 0.647 Fuller’s earth

6 470 0.363

Granular carbon

0.594 Clay 4-40 500 0.102

145

Enhanced Filtration

Filter Performance Evaluation Several measurement techniques could be used to characterize filter performance.

Particle concentrations could be measured using a particle counter, or measured indirectly using a turbidimeter. If the particle suspension absorbed a significant amount of light, a spectrophotometer could be used. A microscope could be used to count particles. If microorganisms are used as the source particles, they could be enumerated using standard microbiological techniques such as membrane filtration followed by growth on selective media.

Turbidimeters measure the amount of light scatter caused by a suspension of particles. Because absorption and scattering of light are influenced by both size and surface characteristics of the suspended material, turbidity is not a direct quantitative measurement of the concentration of suspended solids. In a turbidimeter the scattered light (measured at a right angle to the incident light) and the transmitted light intensities are measured (Figure 2). The ratio of scattered light to transmitted light is proportional to the turbidity of the sample. The constant of proportionality is determined by measuring a known standard.

Experimental Objectives The purpose of this research is to evaluate the ability of the CLSE filter aid to

enhance particle removal in a filter operating at rapid sand filtration rates. We will use tap water amended with kaolin, 2.5 cm diameter filter columns, and turbidimeters. Students will assembly the apparatus.

Experimental Methods 1) Setup 2.5 cm diameter filter column plumbing (Make all connections firmly and

verify that the connections can’t be pulled apart) including 1 L of clay suspension on a stirrer, peristaltic pump for metering in clay suspension and filter aid, flow meter, pressure reducing valve, and pressure sensor for head loss.

2) Add 8 cm of sand to the filter column (by mass). 3) Carefully observe the sand surface as you gradually increase the flow rate from

zero in backwash mode. Measure the pressure required to begin to lift the bed. Continue backwashing the filter to clean the sand until the effluent turbidity is less than 0.5 NTU

90° Detector

Samplecell

Transmittedlight detector

Lamp

Lens Figure 2. Light path in a turbidimeter.

146


4) Obtain head loss (in cm) as a function of flow rate (down flow mode) over a range of 1 to 25 m/hr (8.2 to 204 mL/min) using at least 5 data points. Use the rotometer to measure the flow rate.

5) Challenge the filter with a kaolin suspension (approximately 5 NTU) for 30 minutes to determine baseline filter performance.

6) Backwash the filter 7) Add the filter aid (the amount and method of application will be discussed during

lab) 8) Set the down flow rate to 5 m/hr. 9) Measure the head loss to see if the filter aid increased the head loss 10) Pump a clay suspension into the filter influent so that the influent concentration is

10-mg/L kaolin. Measure effluent turbidity and head loss as a function of time for 30 minutes. Take turbidity measurements every 5 minutes and measure the head loss continuously using the Signal Monitor software.

11) Backwash the filter. 12) If you have time test

the filter again to see if the filter aid improved filter performance even after backwashing.

Figure 3. Picture of experiment setup.

147

Enhanced Filtration

Table 2. Filtration parameters.

parameter symbol value unitsapproach velocity Va 5.0 m/hrcolumn diameter d 2.5 cmcolumn area A 4.9 cm2

Column length Lcolumn 20.0 cmMedia depth L 8.0 cmbulk density of media bulkdensity 1650 kg/m3

mass of media sandmass 64.8 gBackwash velocity Vb 50.0 m/hrflowrate (forward) Qd 40.9 mL/minflowrate (backwash) Qb 409.1 mL/minInfluent clay concentration C0 10.0 mg/Ldilution factor dilution 100clay stock concentration Cconcentrate 1000 mg/Lclay stock flowrate Qc 0.41 mL/minmedia residence time thetam 0.96 mintotal residence time thetac 2.4 min

Prelab Questions 1) How much water is required to operate one of the laboratory filters for 2 hours?

Don’t include the water required to fill the filter initially. 2) Given the dimensions for the filter column, a glass density of 2.65 g/cm3, and

filter porosity of 0.4, estimate the mass of glass beads in one filter column. (Show your calculations.)

3) Draw a plumbing schematic of a filter column that allows you to do the following: Measure the pressure drop across the column using a pressure sensor, reverse the flow of water for backwash, and maintain a high pressure in the filter column to avoid dissolution of gasses.

4) Explain how you will switch the filter from down flow to back wash mode.

Data Analysis 1) Compare the pressure required to begin to lift the bed with the calculated value

based on fluid statics. 2) Plot head loss vs. flow rate for a clean bed and estimate the hydraulic conductivity

of the sand. Is the flow laminar or turbulent? What technique did you use to determine the flow regime?

3) Plot the fraction of influent particles remaining in the effluent vs. time for each run on a single graph.

4) Plot head loss as a function of time for each run on a single graph. 5) Calculate the filter coefficient (equation 12.3) for the filter with and without the

filter aid.

148


Questions for Discussion 1) Did the filter aid make a significant difference in filter performance? 2) How was the head loss affected by the addition of the filter aid? 3) The laboratory filter columns were 8 cm deep. Rapid sand filters have 60 cm of

media. Estimate the fractional bacteria removal for a 60 cm deep filter of media. What assumptions did you make to predict the performance of a 60 cm column?

4) What further experimentation do you recommend?

References Iwasaki, T. 1937. “Some Notes on Sand Filtration” Journal American Water Works

Association 29: 1591. Liljestrand, H. M.; I. M. C. Lo and Y. Shimizu. 1992. “Sorption of humic materials

onto inorganic surfaces for the mitigation of facilitated pollutant transport processes” Proceedings Of The Sixteenth Biennial Conference Of The International Association On Water Pollution Research And Control, Washington, D.C., USA, May 26(1-11): 1221-1228.

Tien, C. and A. Payatakes. 1979. “Advances in Deep Bed Filtration” AIChE Journal 25(5): 737.

Weber-Shirk, M., and R. I. Dick. 1997. Physical-Chemical Mechanisms in Slow Sand Filters. Jour. AWWA. 89:87-100.

Weber-Shirk, M. L. and R. I. Dick (1999). “Bacterivory by a Chrysophyte in Slow Sand Filters.” Water Research 33(3): 631-638.

149

Enhanced Filtration

Lab Prep Notes

Setup

1) Attach two Easy-Load pump heads to the pump drives.

2) Setup turbidimeters and verify that the vials are clean.



magnetic stirrer Fisher Scientific 11-500-7S variable flow digital drive


Easy-Load pump head


Filter columns 100-1095 µl


10-109.5 µl pipette


2100P Turbidimeter

Hach Company 46500-00

2100N Turbidimeter


high pressure flow cell




150


Gas Transfer

Introduction Exchange of gases between aqueous and gaseous phases is an essential element of

many environmental processes. Wastewater treatment plants require enhanced transfer of oxygen into activated sludge tanks to maintain aerobic degradation. Water treatment plants require gas transfer to dissolve chlorine gas or ozone. Gas transfer can also be used to remove unwanted volatile chemicals such as carbon tetrachloride, tetrachloroethylene, trichloroethylene, chloroform, bromdichloromethane, and bromoform from water (Zander et al., 1989). Exchange of a dissolved compound with the atmosphere is controlled by the extent of mixing in the aqueous and gaseous phase, the surface area of the interface, the concentration of the compound in the two phases, and the equilibrium distribution of the compound. Technologies that have been developed to enhance gas transfer include: aeration diffusers, packed-tower air stripping, and membrane stripping. Each of these technologies creates a high interface surface area to enhance gas transfer.

Theory Oxygen transfer is important in many environmental systems. Oxygen transfer is

controlled by the partial pressure of oxygen in the atmosphere (0.21 atm) and the corresponding equilibrium concentration in water (approximately 10 mg/L). According to Henry’s Law, the equilibrium concentration of oxygen in water is proportional to the partial pressure of oxygen in the atmosphere.

Natural bodies of water may be either supersaturated or undersaturated with oxygen depending on the relative magnitude of the sources and sinks of oxygen. Algae can be a significant source of oxygen during active photosynthesis and can produce supersaturation. Algae also deplete oxygen levels during the night.

At high levels of supersaturation dissolved gas will form microbubbles that eventually coalesce, rise, and burst at the water surface. The bubbles provide a very efficient transfer of supersaturated dissolved gas to the gaseous phase, a process that can be observed when the partial pressure of carbon dioxide is decreased by opening a carbonated beverage. Bubble formation by supersaturated gasses also occurs in the environment when cold water in equilibrium with the atmosphere is warmed rapidly. The equilibrium dissolved oxygen concentration is a function of temperature and as the water is warmed the equilibrium concentration decreases (Figure 1).

151

Gas Transfer

Supersaturation of dissolved gases can also occur when water carrying gas bubbles from a water fall or spillway plunges into a deep pool. The pressure increases with depth in the pool and gasses carried deep into the pool dissolve in the water. When the water eventually approaches the surface the pressure decreases and the dissolved gases come out of solution and form bubbles. Bubble formation by supersaturated gases can kill fish (similar to the “bends” in humans) as the bubbles form in the bloodstream.

Gas transfer rate can be modeled as the product of a driving force (the difference between the equilibrium concentration and the actual concentration) and an overall volumetric gas transfer coefficient (a function of the geometry, mixing levels of the system and the solubility of the compound). In equation form

( )*,v̂ l

dCk C C

dt= − 13.1

where C is the dissolved gas concentration, C* is the equilibrium dissolved gas concentration and ,

ˆv lk is the overall volumetric gas transfer coefficient . Although ,

ˆv lk

has dimensions of 1/T, it is a function of the interface surface area (A), the liquid volume (V), the oxygen diffusion coefficient in water (D), and the thickness of the laminar boundary layer (δ) through which the gas must diffuse before the much faster turbulent mixing process can disperse the dissolved gas throughout the reactor.

,ˆ ( , , , )v lk f D A Vδ= 13.2

The overall volumetric gas transfer coefficient is system specific and thus must be evaluated separately for each system of interest (Weber and Digiano, 1996).

A schematic of the gas transfer process is shown in Figure 2. Fickian diffusion controls the gas transfer in the laminar boundary layer. The oxygen concentration in the bulk of the fluid is assumed to be homogeneous due to turbulent mixing and the oxygen concentration above

6789

101112

10 20 30 40

Temperature (°C)

Oxy

gen

(ppm

)

Figure 1. Dissolved oxygen concentrations in equilibrium with the atmosphere as a function of water temperature.

δ

oxygen gas

disso lv edoxygenV/ A

CC*

Figure 2. Single film model of interphase mass transfer of oxygen.

152


the liquid is assumed to be that of the atmosphere. The gas transfer coefficient will increase with the interface area and the diffusion

coefficient and will decrease with the reactor volume and the thickness of the boundary layer. The functional form of the relationship is given by

,ˆ

v l

ADk

V δ= 13.3

Equation 13.1 can be integrated with appropriate initial conditions to obtain the

concentration of oxygen as a function of time. However, care must be taken to ensure that the overall volumetric gas transfer coefficient is not a function of the dissolved oxygen concentration. This dependency can occur where air is pumped through diffusers on the bottom of activated sludge tanks. Rising air bubbles are significantly depleted of oxygen as they rise through the activated sludge tank and the extent of oxygen depletion is a function of the concentration of oxygen in the activated sludge. Integrating equation 13.1 with initial conditions of C = C0 at t = t0

0 0

,*ˆ

C t

v lC t

dCk dt

C C=

−∫ ∫ 13.4

*

, 0*0

ˆln ( )v lC C

k t tC C

−= −

− 13.5

This equation can be linearized so that ,ˆ

v lk is the slope of the line.

( ) ( )* *, 0 , 0

ˆ ˆln lnv l v lC C k t C C k t − = + − − 13.6

The simple gas transfer model given in equation 13.5 is appropriate when the gas transfer coefficient is independent of the dissolved gas concentration. This requirement can be met in systems where the gas bubbles do not change concentration significantly as they rise through the water column. This condition is met when the water column is shallow, the bubbles have large diameters, or the difference between the concentration of dissolved gas and the equilibrium concentration is small.

Experimental Objectives The objectives of this lab are to:

1) illustrate the dependence of gas transfer on gas flow rate. 2) develop a functional relationship between gas flow rate and gas transfer. 3) Explain the theory and use of dissolved oxygen probes. See the appendix of this

manual (page 158) for information on how the dissolved oxygen probe works and how to calibrate it.

A small reactor that meets the conditions of a constant gas transfer coefficient will be used to characterize the dependence of the gas transfer coefficient on the gas flow rate through a simple diffuser. The gas transfer coefficient is a function of the gas

153

Gas Transfer

flow rate because the interface surface area (i.e. the surface area of the air bubbles) increases as the gas flow rate increases.

In order to measure the reaeration rate it is necessary to first remove the oxygen from the reactor. This can be accomplished by bubbling the solution with a gas that contains no oxygen. Nitrogen gas is typically used to remove oxygen from laboratory reactors. Alternately, a reductant can be used. Sulfite is a strong reductant that will reduce dissolved oxygen in the presence of a catalyst.

cobalt2 22 3 4O 2SO 2SO− −+ → 13.7

If complete deoxygenation is desired a 10% excess of sulfite can be added. The sulfite will continue to react with oxygen as oxygen is transferred into the solution. The oxygen concentration can be measured with a dissolved oxygen probe or can be estimated if the temperature is known and equilibrium with the atmosphere assumed (Figure 1).

Experimental Methods The reactor is a 250 mL

polypropylene narrow mouth bottle with an additional port drilled near the neck to allow a hypodermic tubing diffuser to extend into the bottle (Figure 3). The hypodermic tubing diffuser consists of 4 stainless steel hypodermic tubes that are glued to a plug that mates to 1/4” tubing connectors. The hypodermic tubes are bent so that they are spaced at 90° intervals around the perimeter of the bottle. The DO probe funnel/mixer assembly is inserted into the bottle opening. The bottle is placed on a magnetic stirrer and the DO probe is inserted into the funnel/mixer assembly. The spacing of the hypodermic tubing at the perimeter of the bottle is designed to prevent rising bubbles from touching the DO probe membrane. The hypodermic diffuser is connected to a peristaltic pump (or other source of regulated air flow).

1) Calibrate the DO probe (See page 159). 2) Prepare to monitor the dissolved oxygen concentration using the Compumet™

software. Use 5 second data intervals and monitor pH on channel A. The Accumet™ meter and the Compumet™ software think the dissolved oxygen probe is a pH probe. Although the output of the meter is pH, it should be interpreted as mg/L of dissolved oxygen. See page 157 for instructions on using Compumet™ software.

DO probe

Hypodermic diffuser

100 mL beaker Water surface

Stirbar

Figure 3. Apparatus used to measure reaeration rate.

154


3) Add 50 mL of distilled water to a 100 mL beaker. 4) Set the stirrer speed to 5.

5) Add ˜10 mg CoCl2· 6H2O (note this only needs to be added once because it is the catalyst). A stock solution of CoCl2· 6H2O (100 mg/mL – thus add 100 µL) has been prepared to facilitate measurement of small cobalt doses.

6) Set the air flowrate to 50 mL/min (or to the desired flow rate). 7) Turn the air off. 8) Add enough sodium sulfite to deoxygenate the solution. A stock solution of

sodium sulfite (100 mg/mL) has been prepared to facilitate measurement of small sulfite doses. (50 mL of water at 10 mg O2/L = 0.5 mg O2, therefore add 5 mg sodium sulfate or 20 µL of stock solution.)

9) Turn the air on and start collecting data using the Compumet™ software. 10) Monitor the dissolved oxygen concentration until it reaches 80% of saturation

value. 11) Save the data as \\Enviro\enviro\Courses\453\gastran\netid_100 for later analysis.

Repeat steps 6-11 using flow rates of 100, 200, 300, and 500 mL/min.

Prelab Questions 1) Calculate the mass of sodium sulfite needed to reduce all the dissolved oxygen in

50 mL of pure water in equilibrium with the atmosphere and at 30°C. 2) Sketch your expectations for dissolved oxygen concentration as a function of time

for the flow rates used on a single graph. The graph can be done by hand and doesn’t need to have any numbers on the time scale.

3) Sketch your expectations for ,ˆ

v lk as a function of gas flow rate. Do you expect a perfectly straight line or do you expect some nonlinearities? Why? What do you expect ,

ˆv lk to be when the gas flow rate is zero?

Data Analysis 1) Eliminate the data from each data set when the dissolved oxygen concentration

was less than 0.5 mg/L. This will ensure that all of the sulfite has reacted. 2) Set t0 to the time at the beginning of the remaining data. Subtract t0 from each of

the times so the remaining data now starts at zero. 3) Plot the 5 data sets with the corrected times on a single graph.

4) Estimate ,ˆ

v lk using linear regression and equation 13.6 for each data set. Show a graph with the linearized data and the best fit lines.

5) Graph ,ˆ

v lk as a function of gas flow rate.

6) Comment on results and compare with your expectations and with theory.

155

Gas Transfer

References Weber, W. J. J. and F. A. Digiano. 1996. Process Dynamics in Environmental

Systems. New York, John Wiley & Sons, Inc. Zander, A. K.; M. J. Semmens and R. M. Narbaitz. 1989. “Removing VOCs by

membrane stripping” American Water Works Association Journal 81(11): 76-81.

156


Lab Prep Notes

Setup

1) Prepare the sodium sulfite immediately before class and distribute to groups in 15 mL PP bottles to minimize oxygen dissolution and reaction with the sulfite.

2) The cobalt solution can be prepared anytime and stored long term. Distribute to student stations in 15 mL PP bottles.

3) Attach two Easy-Load pump heads to the pump drives and plumb with size 18 tubing joined and connected to the hypodermic diffuser.

4) Verify that DO probes are operational, stable, and can be calibrated. The solution behind the membranes should be clear and free of bubbles. If necessary replace membranes.

5) Mount DO probes on magnetic stirrers.

Table 1. Reagent list Description Supplier Catalog

number Na2SO3 Fisher Scientific S430-500

CoCl2· 6H2O Fisher Scientific C371-100

Table 2. Stock solutions list

reagent M.W. g/100 mL

mg/ mL

mL/ group

solubility g/L

Na2SO3 126.04 10 g 100 10 125 CoCl2· 6H2O

237.92 10 g 100 1 770



magnetic stirrer Fisher Scientific 11-500-7S 100-1095 µL


10-109.5 µL pipette


15 mL PP bottles

Fisher Scientific 02-923-8G



Easy-Load pump head


PharMed tubing # 18


4 prong hypodermic

tubing diffuser

CEE shop

1/4” plug Cole Parmer H-06372-50 1/4” union Cole Parmer H-06372-50

stainless steel hypodermic

tubing

McMaster Carr

157

Instrument Instructions


Compumet software Information on use of the Compumet software is available at

http://www.cee.cornell.edu/mws/Software/Compumet.htm.

pH Probe Calibration

Select U.S. Standard Buffers From the main screen, press the setup key. Setup 1 menu is shown.

1) Press the 1 key to select Buffer Recognition. Observe display of Setup Buffer Recognition.

2) Press the 1 key again to select U.S. standard buffers. 3) Press the enter key to implement the selection. 4) Press the clear key to return to the main screen.

Full Calibration (done once a day) This will calibrate both the slope and the null point. Use either 2 or 3 buffers.

Clear Existing Buffers Press the pH key to select pH measurement. Observe display of the current pH

standardization points. 1) Press the standardization key. A menu of standardization options appears. 2) Press the 2 key to select Clear Existing Standards. 3) The meter returns to the main screen, but with all pH standardization points (for

the current channel) cleared from memory. The electrode slope is reset to 100% or 59.16 mV/pH.

Add Buffers (can use buffers at pH of 4, 7, and 10) 1) Press the standardization key. A menu of standardization options is displayed. 2) Press the 1 key to select Update or Add a Standard. 3) The Prepare Buffer/Standard screen appears. Make sure the electrode is in the

buffer solution. Press the enter key. 4) The meter will wait until an electrode stability criterion is reached. Then it will

automatically read the signal and calibrate. 5) The meter returns to the main screen with the added buffer point shown.

Repeat these steps for each of the buffers.

Updating the Standardization (done every hour) Use a single buffer to adjust the null point of the calibration curve. The calibration

slope will remain the same as from the last full calibration. Press the standardization key. A menu of standardization options is displayed.

158


1) Press the 1 key to select Update or Add a Standard. 2) The Prepare Buffer/Standard screen appears. Make sure the electrode(s) is in the

buffer solution. Press enter. 3) The meter will wait until a pre-determined electrode stability is reached, then it

will automatically read the signal and calibrate.

pH Probe Storage

Preparation for storage

1) Rinse with distilled water 2) Air dry 3) Store for months as needed

Preparation for use

1. Heat in 60°C water and stir for 15 minutes 2. Place in pH buffer 4 or 7 at room temperature for 15 minutes 3. Standardize 4. Probes that fail to standardize may require cleaning

Procedure for Cleaning pH Gel-Filled Polymer Electrode 1. Warm distilled water to 40 - 60 °C

2. Suspend the electrode in the warm water for about 15 min while stirring with a

magnetic stirrer. This will loosen any material attached to the probe

3. Add 1/2 tsp of detergent Terg-A-Zyme∗ to the water. Keep stirring for 15 min

4. Rinse well with distilled water

5. Store probe in pH 4 buffer for at least 3 hr (the longer the better). Although less

preferable, the probe can also be stored in pH 7 buffer

6. Standardize

7. Probes that fail to standardize may need to be discarded

Note: Terg-A-Zyme∗ - Fischer cat. no. 04-322--11A.

Dissolved Oxygen Probe

Theory The probe makes use of the fact that an applied potential of 0.8 V can reduce O2 to

H2O:

159


- +2 24e + 4H + O 2H O→ 13.8

The cell is separated from solution by a gas permeable membrane that allows O2 to pass through (Figure 1). The concentration of O2 in the cell is kept very low by reduction to H2O. The rate at which oxygen diffuses through the gas permeable membrane is proportional to the difference in oxygen concentration across the membrane. The concentration of oxygen in the cell is ˜0 and thus the rate at which oxygen diffuses through the membrane is proportional to the oxygen concentration in the solution.

Oxygen is reduced to water at a silver (Ag) cathode of the probe. Oxygen reduction produces a current that is measured by the meter.

The probe is calibrated using a two-point calibration and can be temperature compensated if desired. Directions can be obtained at http://www.cee.cornell.edu/mws/Software/DOcal.htm.

Monitoring The dissolved oxygen probe output is in

the Pico amp (10-12 A) range. This current must be converted to a voltage in order to be measured by the laboratory data acquisition system. An analog circuit is used to convert the dissolved oxygen probe into a voltage. The voltage is monitored using the Signal Monitor software. Information on using the Signal Monitor software is available at http://www.cee.cornell.edu/mws/Software/signal_monitor.htm.

The 8-pin plug from the signal-conditioning box that is attached to the dissolved oxygen probe needs to be plugged into the middle row of ports on the lab bench.

Calibration

1) A calibration routine is available in the Signal Monitor software. Follow the instructions in the software and use the help as needed. http://www.cee.cornell.edu/mws/Software/DOcal.htm.

Dissolved Oxygen Probe Storage

1) Remove membrane from probe. 2) Rinse electrode with distilled water. 3) Store electrode covered with electrode cap (dry).

probe

KCl electrolyte

A

[O2 (aq)]

solution

Ag (cathode)Ag (anode)

gas permeable membrane

voltage sourcecurrent meter

probe body

Figure 1. Schematic of dissolved oxygen probe.

160


Gas Chromatograph Information on the spectrophotometer analysis software is available at

http://www.cee.cornell.edu/mws/Software/gas_chromatograph.htm.

UV-Vis Spectrophotometer Information on the spectrophotometer analysis software is available at

http://www.cee.cornell.edu/mws/Software/Spectrophotometer.htm.

161

Index

Index

A absorption, 20, 71, 72, 78, 145 acid neutralizing capacity, 43, 45, 46 acid precipitation, 43, 47, 50 adsorption, 24, 70, 71, 72, 74, 75, 82, 83, 128 advective dispersion, 33, 34, 130, 131 alkalinity, 45, 50, 61 ANC, 45, 46, 47, 48, 49, 50, 51, 52, 53, 54, 55, 57,

58, 59, 60, 61, 62, 107, 108, 109 atomic absorption, 78

B BET, 127, 128 Biochemical Oxygen Demand, 88 Biodegradable, 87 Boltzmann constant, 128, 140

C carbonate system, 44, 45, 107 chelating agent, 70, 80 coefficient of variation, 23, 133, 135, 137, 138 completely mixed flow, 30, 39, 53 Compumet, 37, 38, 39, 49, 51, 62, 153, 154, 157 conductivity, 35, 36, 37, 38, 39, 41, 111, 113, 147 correlation coefficient, 21, 24, 28

D density of sodium chloride solution, 25 density of water, 129 deoxygenate, 154 diffusion, 34, 119, 131, 139, 151, 152 diode array, 21, 65, 77 dispersion coefficient, 30, 31, 32, 33, 37, 40, 130 dissolved oxygen, 87, 88, 89, 90, 92, 93, 94, 95, 96,

97, 150, 151, 152, 153, 154, 159

E endogenous respiration, 88, 89, 90, 92, 95, 96 extracellular polymer, 77, 82 extractant, 77, 78, 79, 80, 81, 82, 84

F flow with dispersion, 30, 33

G gas chromatograph, 9, 109, 110, 111, 113, 114, 115,

120, 123, 131, 135, 141 gas transfer, 96, 99, 150, 151, 152 Gas transfer, 150, 151

gasoline, 119 global warming, 87 Gran plot, 53, 58, 59, 61, 62 Gran Plot, 53, 58 groundwater, 33, 119, 130

H hazardous waste site, 70 Henry's Law, 46, 129 hydraulic residence time, 32, 34, 40, 44, 46, 50, 53

I ion exchange, 71, 74 isotherms, 74, 127, 128, 137

L landfill, 86, 100, 115 Langmuir, 128 ligand, 75

M mass balance, 33, 40, 50, 129, 131, 133 methane, 100, 101, 102, 103, 104, 105, 109, 110,

111, 113, 114, 115, 116 methylene blue, 22, 23, 24, 25, 28, 29, 77, 78, 79,

80, 81, 82 Monod, 88, 89

O oxidant, 76 Oxidation, 83, 87, 88 oxygen deficit, 88, 90, 91, 92, 93 oxygen probe, 93, 94, 95, 152, 153, 159 oxygen sag, 88, 93, 96, 97

P Peclet number, 32, 33, 40 petroleum, 119 pH probe, 48, 49, 51, 56, 60, 63, 153 pipette, 11, 23, 55, 61, 66, 69, 95, 149, 156 plug flow, 30, 32, 34, 39 Plug flow, 34 plume, 40 porous media, 33, 37, 40, 41, 55, 56, 70, 71, 73, 74,

75, 76, 77, 119, 129, 130, 131 pressure sensor, 109, 111, 113, 114, 145, 147 pump and treat, 70

162


R reactor, 9, 30, 31, 32, 33, 34, 35, 37, 38, 39, 40, 41,

50, 53, 77, 88, 91, 94, 95, 151, 152, 153 reductant, 153 remediation, 47, 48, 70, 71, 76, 77 Remediation, 43, 47, 49, 53 retardation factor, 130, 131

S Signal Monitor, 146, 159 slow sand filtration, 142 soil density, 122 soil gas sampling, 119, 121 soil moisture content, 122, 123, 129, 132, 137 soil porosity, 138 solvents, 12, 16, 17, 119

sorption, 71, 72, 74, 76, 77, 78, 120, 126, 127, 128, 129, 130, 131, 134, 139

spectrophotometer, 21, 22, 24, 28, 29, 65, 66, 67, 69, 77, 86, 145, 160

spectrum, 21, 66, 78 Streeter Phelps, 88 Superfund, 70

T titration, 52, 57, 58, 59, 60, 61, 62 tracer, 30, 31, 32, 33, 34, 35, 36, 37, 38, 39, 40, 41,

42 turbidimeter, 145 turbidity, 145, 146

W watershed, 43, 46, 47, 48

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