Measuring seagrass photosynthesis: methods and applications

Measuring seagrass photosynthesis: methods andapplications

João Silva1,*, Yoni Sharon2, 3, Rui Santos1, Sven Beer3

1Marine Plant Ecology Research Group, Center of Marine Sciences (CCMar), Universidade do Algarve,Campus de Gambelas, 8005-139 Faro, Portugal

2The Interuniversity Institute for Marine Sciences, POB 469, Eilat 88103, Israel3Department of Plant Sciences, Tel Aviv University, Tel Aviv 69978, Israel

AQUATIC BIOLOGYAquat Biol

Vol. 7: 127–141, 2009doi: 10.3354/ab00173

Printed November 2009Published online October 22, 2009

INTRODUCTION

Early studies on seagrass photosynthesis (e.g. Drew1978) usually began by explaining that seagrasses arerelatively primitive monocotyledons, closely related tofreshwater plants but able to live in the marine envi-ronment. Presently, many introductions start by high-lighting the importance of seagrasses in overallmarine productivity and their subsequent economicalimportance (e.g. Duarte & Cebrián 1996, Costanza etal. 1997, Duarte & Chiscano 1999). This shift of em-phasis indicates the evolution of seagrass perceptionin global biological science and reflects the contribu-tion of the increasing number of physiological andecophysiological studies dealing with this plantgroup.

Seagrass photosynthesis was, until recently, mostcommonly measured in laboratory experiments, usu-ally by incubating leaf segments in closed chambersand determining initial and end O2 values, or by fol-lowing continuous O2 evolution in the chambers withClark-type electrodes. These methods have providedmost of the fundamental information on seagrassresponses to factors such as light, temperature andnutrients (comprehensively reviewed by Lee et al.2007) and on the mechanisms of carbon uptake andfixation by these plants (Beer et al. 2002). However,they are extremely intrusive, as they involve plantremoval from the natural environment and a highdegree of manipulation (Beer et al. 2001).

In situ measurements of photosynthetic activity inseagrasses were made possible after the development

© Inter-Research 2009 · www.int-res.com*Email: [email protected]

ABSTRACT: This review originates from a keynote lecture given at the recent 8th Group for AquaticProductivity (GAP) workshop held in Eilat, Israel. Here we examine the most important methodolo-gies for photosynthetic measurements in seagrasses and evaluate their applications, advantages anddisadvantages, and also point out the most relevant results. The most commonly used methodologiesare based on oxygen (O2) evolution and chlorophyll fluorescence measurements. O2-based method-ologies allowed for the first approaches to evaluate seagrass productivity, whereas chlorophyll a flu-orescence has more recently become the choice method for in situ experiments, particularly in eval-uating photosynthetic responses to light and assessing stress responses. New methodologies havealso emerged, such as O2 optodes, underwater CO2 flux measurements, geo-acoustic inversion andthe eddy correlation technique. However, these new methods still need calibration and validation.Our analysis of the literature also reveals several significant gaps in relevant topics concerning sea-grass photosynthesis, namely the complete absence of studies on deep-growing populations thatphotosynthesise under extreme low light conditions and the uncertainties about the true degree ofseagrass carbon limitation, which limits our ability to predict responses to global changes.

KEY WORDS: Seagrass photosynthesis · O2 evolution · Chlorophyll a fluorescence · Carbon uptake ·CO2 flux

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Contribution to the Theme Section ‘Primary production in seagrasses and microalgae’ OPENPEN ACCESSCCESS

Aquat Biol 7: 127–141, 2009

of a submersible pulse-amplitude modulated (PAM)fluorometer (applied initially by e.g. Beer et al. 1998,Ralph et al. 1998, Björk et al. 1999, Beer & Björk 2000).Recently, an infrared gas analysis (IRGA) techniquehas been adapted for in situ continuous dissolved CO2

flux measurements using incubation chambers con-nected to the analyser at the surface (Silva et al. 2008).While PAM fluorometry determines photosynthetictraits of individual plants, the IRGA method measuresthe rates of gas exchange at the community level. Acombination of both methods may give real-time in-formation on both the photosynthetic characteristics ofseagrass plants and the community CO2 exchange ofundisturbed seagrass meadows.

In situ experiments are less intrusive and deal withthe plants in their natural habitat, hence the photo-synthetic measures incorporate the influences of allambient parameters. However, the control of experi-mental conditions is limited, and insights on specificprocesses are difficult to obtain. On the other hand,laboratory experiments allow for the control andmanipulation of most external parameters and aremost suited to obtain very specific information, e.g.data on biochemical processes related to photosynthe-sis. The general downside of laboratory experiments isthe difficulty in extrapolating results to the naturalenvironment, e.g. time-related patterns of photo-synthetic activity.

We review the fundamental aspects of the most im-portant methodologies used for photosynthetic mea-surements in seagrasses, detailing their field of appli-cation and highlighting the most relevant resultsachieved. Some new and promising techniques arepresented and a critical analysis of the major advan-tages and disadvantages of each method is given. Sig-nificant gaps in this area of research are discussed andpresented as potential pathways for future work.

MEASURING SEAGRASS PHOTOSYNTHESIS

O2 measurements

Measuring the O2 evolved during photosynthesis isone of the oldest and simplest ways of quantifying thephotosynthetic activity of plants. In the aquatic envi-ronment, O2 concentration can be determined: (1)chemically by Winkler titration, (2) polarographicallyusing O2 electrodes, or (3) optically using O2 optodes.

The Winkler method

When applied to seagrasses, the Winkler method (seeStrickland & Parsons 1972, adapted to field use by Drew

& Robertson 1974, Grasshoff et al. 1983) usually involvesenclosing the plants in sealed containers (or ‘bottles’) andleaving them to incubate for a period of time. The in-crease or decrease of the O2 concentration during the in-cubation period provides a measure of either net photo-synthesis (in the illuminated transparent bottles) orrespiration (in darkened bottles), respectively. Grossphotosynthesis can be calculated by correcting the netphotosynthetic rates obtained with the rate of dark (mi-tochondrial) respiration. The Winkler method remainsone of the most accurate ways of measuring dissolved O2

concentrations, arguably better than most commercialfield-going O2 probes. Nevertheless, there are a numberof downsides to this methodology, namely: (1) the needto detach and cut down plant samples (since wholeplants usually do not fit into the bottles), compromisingintegrity and inducing stress (except when this method isused to measure O2 before and after whole-communityincubations); (2) the difficulty in maintaining adequatehomogenization of the medium during incubations, lead-ing to increased boundary layers and resulting in under-estimations of photosynthetic rates (Koch 1994); (3) thelikelihood of O2 saturation and/or inorganic carbon de-pletion during the incubation period, both resulting inphotosynthetic inhibition due to, for example, photo-respiration (Beer 1989) and carbon limitation, respec-tively (neither verifiable in real-time as only initial andfinal O2 concentrations are determined); and (4) the dif-ficulty in ensuring a proper homogeneous or in situ-likeillumination of the plant sample. This method thusallowed for the very first approaches to in situ and labo-ratory experiments of seagrass productivity: Drew(1978, 1979) investigated the seasonal variation in thephotosynthetic activity of Cymodocea nodosa and Posi-donia oceanica, generated photosynthesis-irradiance(P-E) curves and measured dark respiration in C. nodo-sa, Halophila stipulacea, Phyllospadix torreyi, Posidoniaoceanica, Zostera angustifolia and Z. marina, determin-ing the effects of temperature on photosynthesis and cal-culating compensation and saturation irradiances forthose species. Over the years, the effects of light, temper-ature, salinity and pressure on seagrass photosynthesiswere further investigated using this technique in Halo-dule uninervis, Halophila stipulacea and Halophilaovalis (Beer & Waisel 1982, Wahbeh 1983), Z. muelleri(Kerr & Strother 1985), C. nodosa (Pérez & Romero 1992,Zavodnik et al. 1998, Olesen et al. 2002) and Posidoniaoceanica (Olesen et al. 2002). Recently, a few studiesfocusing on whole-community metabolism also usedWinkler titrations to determine O2 concentrations beforeand after incubations in benthic chambers (Barrón et al.2004, Gazeau et al. 2005a,b). Nevertheless, the long in-cubation times used in these studies raise the concernthat photosynthesis may be underestimated due to thedecreased carboxylase activity of Rubisco in response to

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the increase in O2 and decrease in CO2 concentrationswithin the incubation chambers.

O2 electrodes

Electrochemical sensors for O2 measurements arecommonly known as Clark-type O2 electrodes (aftertheir inventor, Leyland C. Clark). These electrodes arecomposed of a platinum cathode and a silver anodeseparated by a salt bridge. An electron flow is set inmotion when a control unit applies a small electricpotential to the system. The current flow is propor-tional to the O2 consumed at the cathode and can beeither analogically plotted or converted into a digitalsignal to be recorded by a control unit (Walker 1987).

Clark-type electrodes are available in a number ofoptions such as handheld units, bench oxymeters orincorporated in complete commercially availablesetups. Whereas handheld units, with low accuracy,are mainly useful for indicative measurements of dis-solved O2 (DO) concentrations, bench-top oxymeters,with better performance, have been used in conjunc-tion with customized incubation chambers, allowingfor versatile and tailor-made apparatuses (Smith et al.1984, Fourqurean & Zieman 1991, Koch 1994, Masiniet al. 1995, Masini & Manning 1997). Complete sys-tems comprised of Clark-type electrodes attached towater-jacketed incubation chambers are also availableon the market, namely those by Hansatech Instru-ments and Rank Brothers. These laboratory systemsprovide the highest available resolution and accuracyand have become standard in photosynthetic research.O2 electrodes such as these are probably among thebest tools to investigate photosynthetic mechanismsand determine the photosynthetic capacities of sea-grasses. Photosynthetic capacity refers strictly to thephotosynthetic rate obtained under ideal conditions,i.e. with non-limiting dissolved inorganic carbon (DIC)supplies, nutrients and light and at optimum tempera-ture (Jones 1994). Such demanding conditions are rel-atively easy to reproduce and maintain in a small,highly controlled environment such as an O2 elec-trode-coupled reaction vessel, but almost impossible toachieve in any other environment.

O2 electrodes are mostly used in 2 general types ofexperimental setups: (1) those in which whole plants orplant sections are incubated for a selected period oftime in a sealed container and O2 concentrations aredetermined at the beginning and at the end of theincubations (endpoint incubations), and (2) those inwhich an O2 electrode is coupled to the incubationchamber and O2 evolution is continuously monitored.

Using endpoint incubations, Evans et al. (1986) eval-uated temperature acclimation effects on the photo-

synthetic rates of Zostera marina and Ruppia maritimaleaf tips (from Chesapeake Bay) while Terrados & Ros(1995) investigated the seasonal variation of tempera-ture effects on the photosynthesis and respiration ofshallow Mediterranean Cymodocea nodosa. Koch &Dawes (1991) searched for differences in P-E curvesbetween 2 ecotypes of R. maritima from Florida andNorth Carolina and Enríquez et al. (1995) determined aseries of P-E relationships in plant sections in anattempt to identify a relationship between the photo-synthetic activity of several Mediterranean macro-phytes and their morphological characteristics. Inverset al. (1997) investigated the effects of pH on photo-synthetic rates of Zostera noltii, C. nodosa and Posido-nia oceanica; Ramírez-García et al. (1998) incubatedpre-desiccated leaves of Phyllospadix scouleri andPhyllospadix torreyi in glass bottles to assess theeffects of different previous air-exposure periods onthe photosynthetic activity; and Ruiz & Romero (2001)compared light response curves among Posidoniaoceanica samples harvested from an in situ multilevelshading experiment. Finally, Alcoverro et al. (1998,2001) examined the seasonal and age-dependence ofPosidonia oceanica photosynthetic parameters and itsannual carbon balance; Invers et al. (1999, 2001) eval-uated the role of carbonic anhydrase in the inorganiccarbon supply to Posidonia oceanica and C. nodosa;and Bintz & Nixon (2001) measured respiration andphotosynthetic rates of whole-plant Z. marina seed-lings at 3 different light levels. Hence this type of setuphas been used to determine photosynthetic lightresponse curves and to evaluate the effects of season-ality, morphological characteristics, leaf age depen-dence, shading, temperature, desiccation and pH onseagrass photosynthesis.

Systems based on continuous O2 measurement havebeen used both in laboratory and field experiments.Most laboratory work has been done using the com-mercially available integrated systems. Dennison &Alberte (1982) investigated the effects of light avail-ability on Zostera marina photosynthesis and growth;seasonal variations in photosynthetic light responseswere evaluated in Z. noltii from the Netherlands (Ver-maat & Verhagen 1996), in Z. marina and Phyllospadixscouleri from California (Cabello-Pasini & Alberte1997) and in 2 Texan Thalassia testudinum populations(Herzka & Dunton 1997); Kaldy & Dunton (1999) deter-mined P-E relationships in T. testudinum laboratory-grown seedlings; Cabello-Pasini et al. (2002) comparedthe maximum photosynthetic rates of an open-oceanZ. marina population with that from a coastal lagoon;Invers et al. (2001) measured continuous O2 evolutionin Z. marina and P. torreyi from the eastern Pacific;Peralta et al. (2000, 2005) compared photosyntheticlight response curves between 2 Z. noltii morphotypes;

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Silva & Santos (2004) correlated photosynthetic O2

release with electron transport rates in the same spe-cies; and Cayabyab & Enríquez (2007) determined thephotosynthetic light responses of T. testudinum plantssubmitted to 3 distinct light treatments. These inte-grated O2 measuring systems have the advantage ofmeasuring net gas exchange, which may be correlativeto growth, but their often small size and the fact thatthe plants (or usually only leaves) must be enclosed,limits the reliability of the results for extrapolation totrue, unobstructed, in situ conditions.

The above-mentioned systems have also beenwidely used in experiments addressing the mecha-nisms of carbon uptake, particularly those concerningthe use of HCO3

– by seagrasses. Using Clark-type O2

electrode-based experimental setups, the use of HCO3–

as a DIC source was demonstrated for Zostera muellerifrom Australia (Millhouse & Strother 1986), Thalassiatestudinum from Florida (Durako 1993), Posidonia aus-tralis from Australia (James & Larkum 1996) and Z.marina (Beer & Rehnberg 1997). Beer & Koch (1996)used this kind of system to simulate primitive inorganiccarbon conditions in the Cretaceous and compare thephotosynthetic performance of Z. marina and T. testu-dinum under those conditions with that under presentday ocean conditions. Zimmerman et al. (1997) evalu-ated the effects of CO2 enrichment on Z. marina pro-ductivity and light requirements with a similar ap-paratus. Björk et al. (1997) investigated inorganiccarbon use in 8 seagrass species from the EasternAfrican coast, by measuring the response of photosyn-thesis to increased inorganic carbon levels. In the samestudy, Halophila ovalis, Cymodocea rotundata andSyringodium isoetifolium also revealed their ability touse HCO3

– as an inorganic carbon source. Hellblom etal. (2001) assessed the effects of commonly usedbuffers on the photosynthetic rates of Z. marina andwere able to identify 2 mechanisms of HCO3

– utiliza-tion, one of them firstly reported for seagrasses basedon extruded protons-mediation of HCO3

– uptake.Mercado et al. (2003) investigated the affinity for DICand the presence of different mechanisms of HCO3

–

use in 2 morphotypes of intertidal Z. noltii from southernPortugal. They found that this species had a low affin-ity for HCO3

–, particularly the lower intertidal morpho-type. Further, photosynthetic pathways, mechanismsof inorganic carbon acquisition, effects of carbonenrichment and buffer additions and the molar rela-tionships between O2 release and electron transporthave been elucidated using such O2 electrode systems.On the other hand, extrapolation of the results ob-tained in these small-volume, laboratory-bound sys-tems to in situ conditions may be rather incorrect.

A few authors have built custom chambers with cou-pled electrodes for specific experiments. Smith et al.

(1984) used a custom-made 2-chambered apparatusconnected to a Clark-type O2 electrode to demonstrateand quantify the transport of O2 from the shoots to theroot-rhizome system of Zostera marina. In a very ele-gant experimental design, the authors incubated intactshoots, separating the leaves from the root-rhizomesystem in 2 adjacent compartments with separatemedia and illumination. By quantifying the O2 takenup or released by the root-rhizome system in theabsence or presence of shoot illumination, it was possi-ble to elucidate the role of the light environment in themaintenance of root aerobiosis. This line of researchwas further pursued by Greve et al. (2003) and Binzeret al. (2005), using microelectrodes inserted directlyinto seagrass rhizomes. These studies significantlyadvanced our understanding of metabolic integrationin seagrasses. They provided the first in situ measure-ments of the fraction of photosynthetically evolved O2

that is channelled down internally to below-ground tis-sues and eventually leaked to ensure aerobiosis in therhizosphere. Fourqurean & Zieman (1991) used a cus-tom-made transparent acrylic chamber with a coupledClark-type polarographic O2 probe to determine thelight-response curves of whole Thalassia testudinumleaves from south Florida. Koch (1994) examined theeffects of hydrodynamics and boundary layer thick-ness on the photosynthetic rates of T. testudinum andCymodocea nodosa. In that study, plant samples wereincubated in a temperature-controlled microcosm withan externally recirculated incubation medium. O2 evo-lution was continuously monitored by a Clark-type O2

electrode installed at an intermediate point of the sys-tem. Masini et al. (1995) and Masini & Manning (1997)determined photosynthetic light responses in Posido-nia sinuosa, P. australis, Amphibolis griffithii and A.antarctica in a custom-built tubular incubation cham-ber with an external water recirculation circuit. Waterflowing in the circuit was passed through a high-resolution O2 sensor, recording O2 values every 15 s.

Some researchers brought high precision O2 elec-trodes to the field, connecting them to benthic incuba-tion chambers to monitor the whole-community O2

release and/or consumption. Dunton & Tomasko (1994)and Major & Dunton (2000) incubated, respectively,whole Halodule wrightii and Syringodium filiformeplants from south Texas, in situ, using transparentacrylic chambers placed on the seabed. Continuous O2

evolution within the chambers was followed using ahigh-resolution dissolved O2 probe installed on a boatat the surface. In both studies, the authors also con-ducted laboratory measurements of P-E curves in leafsegments using water-jacketed incubation chamberswith coupled O2 electrodes. Field and laboratory mea-surements were compared, and in both studies thephotosynthetic efficiency (α), expressed as the initial

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slope of P-E curves, was shown to be highly overesti-mated in laboratory measurements. Other significantdifferences, observed in the compensation intensity (Ic)and in respiration, highlighted the advantages of insitu measurements. Benthic chambers with continuousO2 monitoring were also used by Plus et al. (2001) todetermine primary production and respiration ofZostera noltii beds in the Thau lagoon in South France.

O2 optodes

Optodes are sensors for optical detection of chemicalspecies. The common measuring principle is the use ofan indicator dye (immobilized on the tip of an opticfibre or on a planar surface), the colour of whichchanges as a function of variations in the concentrationof the analyte. Optodes can be used to quantify a num-ber of analytes, e.g. O2 and CO2. Several indicator dyescan be used to build O2 optodes, providing that theirluminescence intensity and lifetime are affected by O2

concentration. A measurement is obtained when anexcitation light is supplied via the fibre and the lumi-nescence quenching of the dye is guided in the oppo-site direction and digitally imaged (Glud 2008). Thequenching magnitude is then directly proportional tothe O2 concentration (Kühl & Polerecky 2008). Op-todes, mostly in their planar configuration, were devel-oped as a tool for 2-dimensional measurements of O2

profiles in benthic communities. The 2-dimensional O2

imaging allows a higher degree of spatial integrationthan single-point microsensor measurements, and thusplanar O2 optodes are seen as a powerful tool to de-scribe spatial and temporal benthic O2 distribution pat-terns and their dynamics (Glud et al. 2001, Glud 2008).

In recent studies (Jensen et al. 2005, Frederiksen &Glud 2006), planar optodes were used for the first timeto investigate O2 leakage in the rhizosphere of the sea-grass Zostera marina. In both cases, O2 imagingrevealed high spatial and temporal dynamics of the O2

distribution in the rhizosphere and identified root tipsas the zone where the oxic areas were more signifi-cant, and thus where most leakage occurred. Oneapparent drawback of planar optodes is that the mea-surements are conducted along an artificial wall,which represents a significant alteration of the naturalsediment structure, with possible effects on O2 spatialdynamics. Tapered O2 micro-optodes provide 1-dimensional measurements and are seen as a possiblealternative to micro-electrodes, with the advantage ofnot consuming O2. Miller & Dunton (2007) essayed theuse of micro-optodes to measure photosynthetic ratesin incubated Laminaria hyperboria discs, and theirresults agreed with published ones obtained by estab-lished methods.

Both planar and tapered O2 optodes appear aspromising techniques for use in seagrass photo-synthetic research. Their full potential is yet to beexplored, and further studies are necessary, particu-larly to compare and validate O2 optodes against otherestablished methods.

Chlorophyll fluorescence

When photons within the photosynthetically activeradiation (PAR) region strike the photosynthetic pig-ment molecules, these become excited. Their excitationenergy is, during the following de-excitation, trans-ferred towards the reaction centre’s special chlorophylla molecules, whose electrons also become excited andare channelled to an electron acceptor molecule(Quinone A, QA). However, a significant portion of theenergy released by de-excitation (or decay) back toground-level does not enter the photochemical processand is, instead, released (or dissipated) as heat or fluo-rescence, i.e. the emission of photons of longer wave-length than the ones absorbed (Schreiber et al. 1998). Itis the nature of an inverse relationship between chloro-phyll a fluorescence and photochemistry that enablesthe use of the former to be used in the investigation ofmany aspects of the photosynthetic process.

Conventional fluorometers emit identical actinic (orphotosynthesis-causing) and excitation lights. Thehigh-wavelength fluorescent light is separated fromthe actinic light by filters before reaching the instru-ment’s detector. This allows for a strong fluorescencesignal, but makes quenching analysis and quantumyield measurements difficult. In modulated fluorome-ters, light is emitted in a succession of alternatelight–dark periods, which enables a more clear sepa-ration of the fluorescence signal from ambient light.PAM fluorometers, the more recent type, emit continu-ous short measuring-light pulses of red or blue light.As the fluorescence signal caused by this measuringlight is captured during the very short pulse periods,external disturbances, background signals and tran-sient artefacts are eliminated and do not mask the flu-orescence signal (Schreiber et al. 1998).

In PAM fluorometers, the short pulses of measuringlight induce the emission of a fluorescence signaltermed Fo or Fs (depending on whether the plant wasdark-adapted or not, respectively). When a saturatinglight pulse of some 0.8 s duration is applied to the plantsample, all reaction centres become reduced (or‘closed’) and the fluorescence emission is maximal (Fm

or Fm’, again dependent on the previous dark-adapta-tion or not, respectively; complete notation of fluores-cence terms can be found in van Kooten & Snel 1990).In the case of dark-adapted plants, the maximum pho-

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tochemical efficiency of PSII is then given by ΦPSII =Fm – Fo/Fm or simply Fv/Fm. This parameter, besidesexpressing the potential for PSII photochemistry, iscommonly used to investigate the onset and recoveryof stress situations due to its sensitivity to plant stress(Beer et al. 2001). In illuminated samples, the samemeasurement provides the effective quantum yield (Y)which is given by Y = Fm’ – Fs/Fm’ = ΔF/Fm’(Genty et al.1989). When Y is measured at steady-state at a knownirradiance (I ), the rate at which electrons are carriedthrough the transport chain past Photosystem II (PSII)(electron transport rate, ETR) can be estimated, ETR =Y × I × AF × 0.5, where AF is the absorption factor (Beeret al. 2001) and 0.5 expresses an equal distribution ofthe absorbed photons between PSII and PSI (Schreiberet al. 1995, see however Grzymski et al. 1997 for alter-native distribution values in some forms of macro-algae). Depending on the research-specific goals, or ifAF cannot be obtained, it may not always be necessaryor relevant to use absolute values of ETR. In suchcases, relative ETRs (rETR), expressed as the productof the effective quantum yield by irradiance, can alsobe used (for a full review of the optical properties andabsorptances of seagrass leaves see Durako 2007).Absolute ETRs are mandatory, for instance, when com-parisons of ETR and gross photosynthetic O2 evolutionare made (Beer et al. 2001). Unlike terrestrial plants,where an AF value of 0.84 is often accepted as a validvalue, seagrass leaves present high variability in thisparameter (Durako 2007).

Chlorophyll fluorescence measurements have beenapplied to seagrass research from the mid-1990s. Twomain types of fluorometers have been used to investi-gate a wide array of questions. A non-modulated orcontinuous excitation fluorometer (PEA, HansatechInstruments) was used in laboratory experiments byDawson & Dennison (1996) to assess the effects ofincreased ultraviolet radiation and elevated PAR onvarious morphological and physiological parameters of5 Australian seagrass species. The same type of instru-ment was also used by Enríquez et al. (2002) to exam-ine variations in photosynthetic activity along theleaves of Thalassia testudinum in the Caribbean Sea.Continuous excitation fluorometers allow for high tem-poral resolution measurements, an essential conditionto investigate the kinetics of the Kautsky inductioncurve (Buschmann & Lichtenthaler 1988, Enríquez etal. 2002). Given that this kind of fluorometry does notallow the separation of fluorescence emission fromambient light, measurements can only be performed indark-adapted samples, which restricts their usefulnessto high quality Fv/Fm determinations and the above-mentioned induction kinetics analysis.

Most research involving fluorescence determinationof seagrasses has been conducted using PAM fluoro-

meters, mainly those produced by Walz. This instru-ment allows measurements to be conducted in full sun-light, thanks to a special emitter-detector unit that sep-arates the fluorescence signal from ambient light(Schreiber et al. 1988). Most PAM fluorometers areportable and one model, the Diving-PAM, is adaptedfor underwater operation. The coupling of these char-acteristics opened the way for autonomous in situmeasurements of effective quantum yields on plantsexposed to natural conditions. Novel, automated, multi-channel chlorophyll fluorometers, first described byRuncie & Riddle (2004) and able to withstand prolongeddeployments, were used to investigate light acclimationprocesses of Halophila stipulacea (Runcie et al. 2009,this Theme Section) during the recent 8th Group forAquatic Productivity (GAP) workshop. Those instru-ments allow autonomous measurements to be con-ducted for longer periods (>24 h), opening new and ex-citing possibilities in seagrass photosynthetic research.Whether in the field or in laboratory experiments, PAMfluorescence has been used in the evaluation of time- orspace-related variations in photosynthesis, with in-sights on the dynamic behaviour of the photosyntheticapparatus of several seagrass species (Silva & Santos2003). P-E curves can be obtained by PAM fluorescencewhen Y is determined along an irradiance gradient. Ifthis gradient is imposed by the fluorometer’s own ac-tinic light source, usually only short (up to minutes)light steps are allowed, due to instrument limitations.The resulting curves are named rapid light curves(RLCs) due to this feature, and must be interpreted dif-ferently from conventional light response curves, al-though they are graphically similar. The critical differ-ence between RLCs and conventional P-E curves is thatthe ETRs used to draw the curves are not measured atsteady-state conditions, given the short exposure ateach light step (Ralph & Gademann 2005 and refer-ences therein). Still, RLCs provide important ecophysi-ological information, namely through the interpretationof curve defining parameters, similar to those of con-ventional light response curves (Beer et al. 1998, Ralph& Gademann 2005, Saroussi & Beer 2007).

Conventional light response curves can be deter-mined in nature using chlorophyll fluorescence, aslong as an external light source supplies moreextended periods of illumination at each light step.This type of curve has been obtained mainly with thegoal of comparing ETRs with gross O2 production inthe search for an expedient and field-going methodol-ogy to estimate photosynthetic rates. In these experi-ments, O2 production and ETRs are simultaneouslymeasured at several irradiances, always at steadystate. Considering that 4 mol of electrons are carriedthrough the transport chain for each mol of O2 pro-duced, there is a theoretical molar ratio of 0.25

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between O2 production and ETR (Walker 1987). Inorder for ETR measurements to be used as proxies forestimations of actual photosynthetic production, 2basic conditions must be met: both a linear relationshipbetween O2 production and ETR, and a 0.25 O2/ETRmolar ratio must be observed, preferably along a com-prehensive range of light levels (Silva & Santos 2004).

Beer et al. (1998) published the first comparison be-tween photosynthetic O2 evolution and ETRs in sea-grasses. The authors examined the O2/ETR relation-ship in Cymodocea nodosa, Halophila stipulacea andZostera marina and found molar values of 0.3, 0.12 and0.5, respectively, although only C. nodosa presented aclearly linear relationship between the evolved O2 andthe transported electrons. Beer & Björk (2000) ob-served a linear relationship and a ratio of 0.28 in Halo-phila ovalis, and a contrasting curvilinear relationshipand a ratio of 0.57 in Halodule wrightii. Silva & Santos(2004) obtained a linear relationship and a ratio of 0.15in Zostera noltii. Enríquez & Rodríguez-Román (2006)verified a linear relationship and a ratio of 0.25 for Tha-lassia testudinum, but only for irradiances <170 µmolm–2s–1. Common to all these studies are: (1) the obser-vation of quite disparate molar ratios, many of them farfrom the theoretical stoichiometry and ranging from0.12 to 0.57, (2) the recording of different initial slopesof ETR and O2 response curves, and (3) the fact thatETR tends to saturate at higher irradiances than O2

evolution. Putative explanations for problems concern-ing ETR calculations are electron-cycling around PSIand non-photochemical quenching in PSII reactioncentres at saturating light levels (Franklin & Badger2001). Uncertainties in O2 evolution may be related tophotorespiration, which can be an important O2 sink(e.g. Flexas & Medrano 2002 and references therein),or the Mehler reaction, in which O2 is photoreduced atPSI with the production of superoxide radicals (Asada1999). The Mehler reaction alone can represent up to10% of the ETR (Makino et al. 2002), corresponding toa similar deviation in the theoretical stoichiometry ofthe photosynthetic process. The conversion of net togross photosynthesis is also likely to introduce addi-tional errors (discussed in Silva & Santos 2004), de-pending on how dark respiration is estimated.

One of the most common applications of PAM fluo-rescence in seagrass research has been the study oftemporal and spatial variation in photosynthesis pat-terns. In a set of laboratory experiments, Ralph (1996)examined the effects of daily irradiance patterns in thephotosynthetic activity of Halophila ovalis, comparinglaboratory-grown plants with wild ones, and observedvery different patterns among the 2 groups in responseto different light regimes. Since these experiments, andtaking advantage of the portability and water resis-tance of the new Diving-PAM, most of the studies ad-

dressing similar questions have been conducted in situ,either in the intertidal area or underwater, usingSCUBA. Daily variation in seagrass photosynthesis, asinfluenced by ambient irradiance, has been investi-gated in situ for Zostera noltii and Cymodocea nodosain southern Europe (Silva & Santos 2003), Posidoniaaustralis in Australia (Runcie & Durako 2004), Halo-phila stipulacea in the Red Sea (Sharon & Beer 2008)and Thalassia testudinum in Florida (Belshe et al. 2008).Spatial variation in photosynthetic activity has alsobeen investigated, at scales ranging from within- toamong-shoot variability up to landscape-level patterns.Ralph & Gademann (1999) measured Fv/Fm ratios alongthe leaves of Posidonia australis while assessing the ef-fect of epiphytes on the plants’ photosynthesis. Durako& Kunzelman (2002) evaluated the variability of photo-synthesis within the shoots of T. testudinum in Florida,and went further, examining differences betweenshoots from healthy and die-off patches and looking attemporal variation among ca. 300 sampling stations dis-tributed among several basins. Turner & Schwarz(2006) conducted a similar study on Z. capricorni fromNew Zealand. Cayabyab & Enríquez (2007) evaluatedthe photoacclimatory responses of T. testudinum plantsto 3 distinct light treatments, comparing the daily varia-tion in photochemical efficiency between basal andapical leaf sections.

Light availability is often regarded as the most impor-tant single parameter controlling seagrass distribution.Responses to light have been widely explored by manyauthors and from a number of different perspectives. Afew studies have particularly addressed the relation-ships between photosynthesis and both depth- andturbidity-related light attenuation. Using the Diving-PAM, Ralph et al. (1998) measured the in situ photo-synthetic responses of Posidonia australis, P. sinuosa,Amphibolis antarctica, A. griffithii and Halophila ovalisfrom shallow Australian waters (0 to 6 m), comparingthem with laboratory measurements of the deepergrowing P. angustifolia (27 m) and Thalassodendronpachyrhizu (46 m). Despite the use of 2 contrasting ex-perimental approaches, the authors found significantlydifferent patterns of photosynthetic behavior betweenshallow and deeper growing seagrasses. Schwarz &Hellblom (2002) measured the photosynthetic light re-sponses of Halophila stipulacea growing at differentdepths in the Red Sea. They found patterns of acclima-tion to distinct light environments, even though theyonly sampled a small depth range (7 to 30 m from aknown 0 to 70 m total range). The same approach wasused by Durako et al. (2003) to compare the photobiol-ogy of Halophila johnsonii and H. decipiens along adepth gradient in Florida, with the goal of explainingtheir relative depth distributions. Additionally, Collieret al. (2008) used Diving-PAM measurements to com-

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plement their physiological characterization of Aus-tralian Posidonia sinuosa along a very small verticalgradient down to 9 m depth. Comparing a broaderspectrum of irradiance controlling factors, Campbell etal. (2003, 2007) used PAM fluorescence to evaluate thecombination of light with other environmental factorson the distribution of several tropical seagrass species,comparing 4 major Australian habitats (estuarine,coastal, reef and deepwater).

Chlorophyll fluorescence has provided detailed de-scriptions of variation in seagrass photosynthetic per-formance at a number of spatial and temporal scales,essentially through the analysis of the quantum effi-ciency fluctuations. Photosynthetic responses to lighthave also been elucidated across a wide range of nat-ural conditions of light quantity and quality.

PAM fluorescence has also been used to investigateinorganic carbon limitations to seagrass photosyn-thesis. Schwarz et al. (2000) used a Diving-PAM con-nected to a small chamber to evaluate in situ the in-organic carbon limitation to Halophila ovalis andCymodocea serrulata photosynthesis. The underwateroperating chamber was used to hold and isolate still-attached leaves, while permitting the addition of inor-ganic carbon, buffers or inhibitors to the leaf-adjacentmedium. PAM fluorescence was also used by Enríquez& Rodríguez-Román (2006) to analyze the photo-synthetic response of Thalassia testudinum to carbonlimitation, as influenced by water flow, in a set of labo-ratory experiments.

Chlorophyll fluorescence has proven to be a veryuseful tool in assessing the onset and recovery ofnumerous types of stress on seagrasses. As discussedabove, the potential quantum yield of PSII, Fv/Fm, is avery useful indicator of stress conditions, and thereforeis widely used in studies addressing a wide variety ofstress situations. Desiccation is a major stressor inintertidal seagrasses. Hanelt et al. (1994) investigatedlow-tide stress in Thalassia hemprichii in South Chinaand observed a midday depression in Fv/Fm, particu-larly when low tide occurred around solar noon, whendesiccation effects were enhanced by high irradiances.The desiccation tolerance of several tropical intertidalseagrasses was evaluated by Björk et al. (1999) inZanzibar. Interestingly, in that study, shallow intertidalspecies were overall more resistant to desiccation thandeeper growing ones, leading the authors to concludethat desiccation tolerance is not the key factor in deter-mining the vertical positioning of seagrass species inthe intertidal zone, but rather their ability to withstandhigh irradiances. In a subsequent investigation inZanzibar, Beer et al. (2006) compared the photo-synthetic responses of H. ovalis, Cymodocea rotundataand Thalassia hemprichii in low-tide conditions, to findthat the first species can only survive in monospecific

tidal pools, given that the presence of the other 2 plantsraises the pH above its compensation point.

The combined effects of desiccation and high tem-peratures were compared in Australian Posidonia aus-tralis and Amphibolis antarctica by Seddon & Cheshire(2001). Thalassia hemprichii and Halodule uninervisfrom Taiwan were also compared by Lan et al. (2005)regarding their relative capacity to deal with both highirradiances and air exposure. The stress responses tohigh irradiance regimes have been mostly investigatedin laboratory experiments, under well-controlled con-ditions, namely for Halophila ovalis (Ralph & Burchett1995, Ralph 1999a) and Zostera marina (Ralph et al.2002). Figueroa et al. (2002) and Kunzelman et al.(2005) approached UV radiation stress by evaluatingthe responses of Posidonia oceanica and Halophilajohnsonii to combinations of PAR with UV-A and/orUV-B, respectively. Other types of stress, whose effectshave been evaluated through the PAM fluorescencetechnique, include thermal stress (Halophila ovalis,Ralph 1998; H. ovalis, Zostera capricorni, Syringodiumisoetifolium, Campbell et al. 2006), osmotic stress (H.ovalis, Ralph 1998), herbicide toxicity (H. ovalis, Z.capricorni, Cymodocea serrulata, Haynes et al. 2000;H. ovalis, Ralph 2000), ammonium toxicity (Brun et al.2008) or even a combination of factors such as temper-ature, light and salinity (Ralph 1999b).

Overall, PAM fluorometry has the advantage ofbeing quick and non-intrusive, and since no enclo-sures are needed, it is particularly suited for in situmeasurements. On the other hand, since respiration isignored in the measurements (as only photosynthesisper se is measured), it is often impossible, or at leastdifficult, to compare such measurements with growthrates. Indeed, the most suitable questions to ask usingPAM fluorometry are those concerning the photo-synthetic light responses to the whole range of ambientparameters.

Dissolved inorganic carbon uptake and CO2 fluxes

Gas molecules, with the exception of those com-posed by 2 identical atoms, absorb radiation at ex-tremely narrow bandwidths within the infrared regionof the spectrum. CO2 has its main absorption peak atλ = 4.25 µm, with 3 secondary peaks at 2.66, 2.77 and14.99 µm (Long & Hallgren 1985). Infrared gas analysis(IRGA) has therefore long been used to measure, withhigh accuracy, the evolution of CO2 exchanged ineither the photosynthetic or respiratory process in ter-restrial plants. A number of laboratory and portablegas analyzers are available, with varying configura-tions, from closed to semi-closed or even open systems,depending on the type of enclosure and air pathway

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(Long & Hallgren 1985). Compared to the previouslydiscussed methodologies, direct assessments of DICuptake or CO2 flux measurements have rarely beenused to estimate seagrass productivity; 4 distinct typesof approaches have been described.

(1) Direct measurement of CO2 uptake by individualleaves, which are enclosed in a mini-cuvette with tem-perature and humidity control coupled to a portableinfrared gas analyzer. Given the considerable exten-sions of intertidal seagrass meadows worldwide, andthe array of information likely to be obtained in situ bythese gas-exchange instruments (to which PAM fluo-rometry can also be coupled), it is surprising that onlya few studies have used this approach (Leuschner &Rees 1993, Leuschner et al. 1998, both with Zosteramarina and Z. noltii). These highly controlled andaccurate systems are, however, limited to intertidalhabitats, as no adaptations are yet made to operatethem underwater.

(2) DIC uptake at the community level has beenmostly investigated using benthic incubations inclosed chambers, where water is retained for a fewhours. Initial and final DIC values are derived fromvariations in pH and alkalinity. As long incubationtimes are required for pH and alkalinity variations tooccur, photosynthesis tends to saturate as Rubisco car-boxylase activity decreases in response to the increas-ing O2 and decreasing CO2 concentrations within thechambers. No assessment has yet been made of howunderestimated the carbon uptake measurements maybe when using this technique. Using both transparentand dark chambers, the net community production anddark respiration are measured, providing an overallgross community production value. Studies based onthis approach have been conducted on Zostera marina(Ibarra-Obando et al. 2004, Martin et al. 2005) andPosidonia oceanica communities (Barrón et al. 2006),complemented with O2 measurements.

(3) Silva et al. (2005, 2008) proposed a variation tothe benthic incubation method by introducing waterrecirculation in the chambers and reducing the lengthof the incubations to minutes instead of hours. In thissystem, water from the chambers is recirculated by aperistaltic pump at the surface and flows through anequilibrator, which allows the partial pressure of CO2

(pCO2) to be continuously monitored in the gas phaseby an infrared gas analyzer. It is important that pH andalkalinity are measured before and after the incuba-tion period, so as to provide an estimate of the total DICflux. This method may also underestimate seagrasscommunity production as CO2 uptake is being regen-erated through carbonate equilibrium dynamics. Fur-ther discussion on the technical and analytical aspectsof this method can be found in Abril (2009) and Silva &Santos (2009).

(4) Air–sea CO2 fluxes have also been used to esti-mate the net community production of seagrass-dominated coastal systems (Gazeau et al. 2005b). Theair–sea CO2 flux is computed from the air–sea gradi-ent of pCO2, the gas transfer velocity and the solubilitycoefficient of CO2. These fluxes can be determinedeither by following pCO2 evolution inside a floatingbell system (Frankignoulle & Distèche 1984) or bysimultaneous measurements of both air and waterpCO2 (Gazeau et al. 2005a, 2005b).

Other methods

Radioactively labelled carbon (14CO2 or H14CO3–)

was one of the first (Beer et al. 2001) methods to deter-mine CO2 uptake. The 14C technique has mostly beenused to investigate the mechanisms of carbon uptakein seagrasses (Beer & Waisel 1979, Abel 1984) and itsproductivity (Williams & McRoy 1982), but nowadays itis only rarely used.

New methodological possibilities are emerging, par-ticularly those addressing large-scale evaluations ofseagrass communitiy production. Hermand et al.(2001) and Hermand (2006) summarized the applica-bility of geo-acoustic inversion techniques to monitorphotosynthetic O2 release in Posidonia oceanica mead-ows. The working principle, verified in this technique,was that the presence of non-dissolved gases in theleaves’ aerenchyma and the production of O2 micro-bubbles sticking to the blade surface interfere with theimpulse response of broad-band acoustic transmis-sions and can be correlated with whole-meadowphotosynthetic activity. This method may provide con-tinuous measurements of oxygen produced by sea-grasses, and may be useful in monitoring seagrass pro-ductivity in large coastal areas. However, moredevelopments of this technique, including calibrationwith more established methods and extrapolation todifferent seagrass species, are necessary.

The eddy correlation technique, used in terrestrialecosystems, was adapted to aquatic environments byBerg et al. (2003). This technique determines the sedi-ment–water fluxes of dissolved O2. The general oper-ating principle lies in the simultaneous measurementof vertical water column velocity and O2 concen-tration at a point a given distance from the sedimentsurface (Berg & Huettel 2008). This is a completelynon-intrusive approach, capable of integrating con-siderable sediment areas (Berg et al. 2007). Althoughthe technique’s full potential is yet to be explored, itappears very promising, particularly in heterogeneousenvironments where its integrating capacity is anadvantage. If used above seagrass meadows, thismethod may provide whole-ecosystem metabolism

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information. The first attempt to determine oxygenfluxes and ecosystem metabolism in seagrasses wasmade by Hume et al. (unpubl. data) in a Zosteramarina community.

CONCLUSIONS

Table 1 summarizes the applications, advantagesand disadvantages of the methods presented above tomeasure seagrass photosynthesis and production. Inroughly 3 decades of research on seagrass photo-synthesis, much information has been gained at differ-ent levels through the use of the various methodologiesdescribed here. O2 measurements, either by Winklertitration or by electrodes, provide accurate rates of netphotosynthetic gas exchange in the light and respira-tion in darkness. However, limitations include the needto use enclosures which, invariably, alter natural irra-diances and water flow conditions. O2 electrodes arealso usually employed in the laboratory, with obviouslimitations related to the removal of plants from theirnatural environments. Measurements of CO2 employ-ing IRGA are much more sensitive than O2 measure-ments and thus require lower incubation times. Whilethe IRGA technique is probably the best for measure-ments of intertidal seagrasses exposed to the air, itsrecent adaptation for measuring dissolved CO2 fluxesof underwater communities may underestimate them.Currently, the most employed method for in situ photo-synthetic measurements is PAM fluorometry. It is fast(a measure of ETR can be obtained within 1 s), and noenclosures are necessary. One major restriction is thatrespiration rates cannot be obtained by PAM fluorom-etry, and photosynthetic rates thus cannot be readilyconverted to production rates. However, this method issuperb for investigating the stresses that impede pho-tosynthesis (by measuring Fv/Fm) as well as for estimat-ing photosynthetic responses to irradiance (includingthe use of RLCs). In the future, it is desirable thatautonomous instruments that measure both chloro-phyll fluorescence and CO2 gas exchange becomeavailable, so that many of the relevant seagrass photo-synthetic parameters can be measured simultaneouslyover extended time periods.

Being rooted plants, seagrasses present the mostcomplex physical structure among marine autotrophs,as plants simultaneously occupy 2 distinct environ-ments, the water column and the sediment. The inte-grated physiology of above- and below-ground tissuesadds a considerable degree of complexity to metabolicstudies, namely in the accurate determination of CO2

fixation and O2 production rates. In fact, although it may represent more than half of the whole-plant O2 consumption (Fourqurean & Zieman 1991), root-

rhizome respiration is rather difficult to measure insitu, and highly unrealistic if measured in laboratoryconditions. On the other hand, a significant portion ofthe photosynthetically evolved O2 is conveyed downfrom the leaves to the below-ground tissues in order tosupport respiration and also to maintain some degreeof aerobiosis in the rhizosphere, which is often sur-rounded by anoxic sediment (Larkum et al. 1989). Inthis context, despite several laboratory studies (Smithet al. 1984, Connell et al. 1999), it was Greve et al.(2003) and Binzer et al. (2005) who firstly providedaccurate in situ measurements of O2 leakage by sea-grass roots and rhizomes, using O2 micro-electrodes.O2 optodes (Glud 2008) added a second dimension tothese measurements. Whereas respiration measure-ments at the plant level are fairly simple in the labora-tory, their extension to the community level has lowvalue. On the other hand, community measurementsstill present a great number of uncertainty factors (out-lined in Middelburg et al. 2005). One of the paths toexplore further in seagrass photosynthetic productionresearch is the assessment of the metabolic contribu-tion of other autotrophic and heterotrophic compo-nents of seagrass communities. Process discriminationand separate metabolic evaluations are essential, inparticular sediment respiration, to avoid consideringthe sediment as just a black box within the community.This will help obtain accurate metabolic budgets forseagrass communities, thus closing the gap betweengross and net primary production.

This review of the published literature on seagrassphotosynthesis, although oriented to methodologicalaspects, also provided an assessment of the status ofseagrass photosynthesis research. We agree with arecent review of the impact of light limitation on sea-grasses (Ralph et al. 2007): although a great deal isknown on seagrass ecophysiology, much information isstill missing, including many fundamental photo-biological data. Our analysis of the literature revealed,for example, that even though seagrasses have a widedepth distribution gradient, down to 50–70 m (denHartog 1970), hardly any work has been done on deeppopulations. This is a major gap in knowledge, espe-cially given the importance of understanding the biol-ogy of plants living in extreme environments and tak-ing into account that seagrass declines worldwide areattributed largely to reductions in light availability(Ralph et al. 2007 and references therein). The avail-ability of underwater instruments such as the Diving-PAM, and the fact that such deep populations arewithin the depth range for technical diving, providesan opportunity for narrowing these gaps in knowl-edge.

With the development of the new large-scale assess-ment techniques described above, research questions

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Method

O2 titration (Winkler)

O2 electrodes coupledto small reactionchambers

O2 microelectrodes

O2 optodes

PAM fluorescence

CO2 evolution

Geo-acoustics

Eddy correlation

Applications

• Photosynthesis and darkrespiration of whole plantsor leaf cuts incubated inbottles (laboratory or insitu)

• O2 analysis of watersamples from benthicchambers (field)

• Photosynthesis and darkrespiration of leaf cuts(laboratory)

• O2 consumption by below-ground tissues (in situ)

• O2 leakage into the rhizo-sphere (in situ)

• O2 production or con-sumption in custom-madechambers

• Used in the eddy correla-tion technique (in situ)

• O2 mapping of seagrassrhizosphere (in situ)

• In situ and laboratorymeasurements of photo-synthetic efficiency at theplant level

• In situ measurements ofcommunity uptake andrelease of CO2, in incu-bation chambers

• In situ large-scale estima-tion of community O2 prod-uction

• In situ sediment–waterfluxes of dissolved O2

• Community level O2

fluxes metabolic studies

Advantages

• High accuracy• Low price

• High resolution• High accuracy• Continuous O2 measure-

ments• Highly controlled condi-

tions• Possibility to manipulate

the incubation medium

• Not very intrusive (smalldiameter electrodes)

• Fast response time• Positioning possibilities

• Not very intrusive• 2-dimensional measure-

ments• Very sensitive at low O2

concentrations• Does not consume O2

• Long-term stability

• Non-intrusive• Portability• Autonomous underwater

equipment• Possibility of continuous

measurements

• Non-intrusive• Integration of whole-

community metabolism• Highly reliable in air-

exposed conditions

• Large-scale application,suitable for ecosystemlevel studies

• Continuous measurements

• Non-intrusive• Autonomous underwater

equipment• Good surface integrating

capacity• Continuous measurements

Disadvantages

• Intrusive (if plants areincubated in bottles)

• Problems related tocontainment in closedchambers

• Initial and final O2

concentrations only• Cumbersome

• Intrusive• Highly artificial• Spectral quality of arti-

ficial light sources

• Small spatial resolution• Fragile in field condi-

tions

• Slower response thanmicroelectrodes

• Technique still underdevelopment

• Measures light reactionsonly

• Does not allow respira-tion measurements orthus production esti-mates

• Possibility of underesti-mating CO2 uptake inunderwater conditions

• Problems related tocontainment in closedchambers

• Underdevelopedtechnique

• Underdevelopedtechnique

Table 1. Applications, advantages and disadvantages of the most common methods used in seagrass photosynthesis and communitymetabolism studies

Aquat Biol 7: 127–141, 2009

pertaining to seagrass meadows as global CO2 sinksmay be answered in the coming years. The effects ofglobal warming on seagrass metabolism also requirefurther research, particularly concerning the effects ofrespiration. Additionally, ocean acidification with theconcomitant CO2 increase is expected to positivelyaffect seagrass photosynthetic rates, but experimentalstudies are scarce (Palacios & Zimmermann 2007).Another interesting aspect to explore related to oceanacidification is the interaction between seagrass photo-synthesis and the calcification rates of other organismswithin the community (e.g. Semesi et al. 2009). Wethus expect future research to bridge the gap betweenthe photosynthetic behaviour of individual plants andthe various aspects of community-level metabolism.

Acknowledgements. This review was presented as a keynotelecture at the 8th International Workshop of the Group forAquatic Primary Productivity (GAP) and Batsheva de Roth-schild Seminar on Gross and Net Primary Productivity, held atthe Interuniversity Institute for Marine Sciences, Eilat, Israel,in April 2008. We thank the Batsheva de Rothschild Founda-tion, Bar Ilan University, the Moshe Shilo Center for MarineBiogeochemistry and the staff of the Interuniversity Institutefor funding and logistic support. J.S. thanks the organizers ofthe workshop, in particular I. Berman-Frank, for the opportu-nity to give this talk and for the wonderful welcome in Eilat.

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Submitted: December 29, 2008; Accepted: June 19, 2009 Proofs received from author(s): September 15, 2009