Penggunaan mencit sebagai hewan
laboratorium disebabkan beberapa sifat
yang menguntungkan:
Mudah dipelihara dalam jumlah besar
Menghasilkan banyak keturunan (6-15 ekor)
Masa reproduktif aktif yang relatif panjang (2-14 bulan)
Masa kehamilan yang relatif pendek
Berat badan mencit dewasa:
Jantan : 20--40 g
Betina : 18--35 g
Mencit dapat hidup dalam jangka waktu:
1 sampai 3 tahun
Pubertas mencit jantan :
Ditandai dengan turunnya testis ke
kantung skrotum . Umur 40 hari
Mencit jantan mencapai kesuburan maks:
Umur 100--300 hari
Mencit betina mempunyai siklus berahi
Poliestrus, dengan tahapan sbb:
- Diestrus
- Proestrus
- Estrus
- Metestrus
Lama satu siklus:
4--5 hari
Tahapan siklus yang sedang berlangsung
dapat diketahui dengan:
Metode Oles Vagina
Lama hidup 1--2 tahun, bisa sampai 3
tahun
Lama produksi ekonomis 9 bulan
Lama bunting 19--21 hari
Kawin sesudah beranak 1 sampai 24 jam
Umur disapih 21 hari
Umur dewasa 35 hari
Umur dikawinkan 8 minggu (jantan dan betina)
Siklus kelamin Poliestrus
Lama siklus estrus 4--5 hari
Lama masa estrus 12--14 jam
Perkawinan Pada waktu estrus
Ovulasi Dekat akhir estrus, spontan
Fertilisasi 2 jam sesudah kawin
Cleavage, zigot blastula 2,5--4 hari
Implantasi 4--5 hari sesudah fertilisasi
Berat dewasa 20--40 g
Berat lahir 0,5--1,0 g
Jumlah anak ~6, dapat mencapai 15
Suhu (rektal) 36--39o C
Pernapasan 140--180/menit; anestesi
80/menit; stres 230/menit
Denyut jantung 600--650/menit; anestesi
350/menit; stres 750/menit
Tekanan darah 130--160 sistol, 102--110 diastol;
anestesi 110 sitol, 80 diastol
Putting susu 10 putting (3 pasang di daerah
dada, 2 pasang di daerah perut)
Plasenta Diskoidal hemokorial
Uterus 2 kornu, bermuara sebelum
serviks
Perkawinan kelompok 4 betina dengan 1 jantan
Kromosom 2n = 40
Aktivitas Nokturnal
Mencit laboratorium dapat ditempatkan dalam kandang yang tidak besar (sebesar kotak sepatu)
Kotak tersebut dapat terbuat dari:
- Plastik
- Aluminium
- Baja tahan karat
Bahan harus tahan lama, dan tahan terhadap gigitan mencit.
Table 1.0.1 Minimum Space Requirements for Housing Laboratory Animals
in Cagesa
Weight Floor area/animal Cage heightb
Animal (g) (in2) (cm2) (in) (cm)
Mouse 25 15.0 96.78 5 12.70
Rat 500 70.0 451.64 7 17.78
a Guidelines are derived from Guide for the Care and Use of Laboratory Animals (ILAR, 1985).b From the resting floor to the cage top From Current Protocols in Immunology Online
Minimizing Risk =
Minimizing Exposure
Procedures to Reduce
Exposure
INDIVIDUALLY VENTILATED CAGES
laminar air rack
From TENCIPLAST
From TENCIPLAST
From TENCIPLAST
Ventilation efficiency with low velocity Air
SpeedVisibility
Easy access
Diuresis cage
From TENCIPLAST
Metabolic cage
From TENCIPLAST
Lingkungan harus kering dan bersih
Suhu sekitar 25oC
Memberi ruangan yang cukup untuk bergerak bebas dalam berbagai posisi
Harus dilengkapi dengan makanan dan minuman yang mudah dicapai oleh
mencit.
Kandang harus ditempatkan dalam rak yang cukup longgar
Kandang harus memiliki alas tidur yang bersih dan berkualitas baik
Table 1.0.2 Recommended Relative Humidity and Dry Bulb
Temperature for Animals Housed in Cagesa
Dry bulb temperatureb
Animal Relative humidity (%)
C F
Mouse 40-70 18-26 64.4-78.8
Rat 40-70 18-26 64.4-78.8
Hamster 40-70 18-26 64.4-78.8
Rabbit 40-60 16-21 60.8-69.8
a Guidelines from ILAR, 1985.
b ILAR, 1965, 1977.
From Current Protocols in Immunology
Harus tidak menarik mencit untuk dimakan
Tidak menarik binatang lain (Misal kutu)
Harus dapat mengisap air
Tidak mengandung zat-zat yang dapat mengganggu penelitian
Alas tidur harus diganti minimal 2 kali dalam seminggu atau bila sudah tercium bau amonia
Alas tidur yang biasa digunakan:
Serbuk gergaji yang tidak terlalu berdebu
Sekam padi yang tidak tercemar oleh kencing ataupun tinja mencit atau tikus liar
Makanan berbentuk pelet dalam jumlah tanpa batas (ad libitum)
Makanan ditempatkan dalam tutup kandang yang
melekuk
Seekor mencit dewasa makan sekitar 3-5 g/h
Kandungan bahan makanan:
- Protein 20-25%
- Lemak 10-12%
- Karbohidrat 40-55%
- Serat kasar ~ 4%
- Abu 5-6%
- Vitamin A 15.000-20.000 IU/kg
- Vitamin D 5000 IU/kg
- Alfa tokoferol 50 mg/kg
- Asam linoleat 5-10 g/kg
- Tiamin 15-20 mg/kg
- Riboflain 8 mg/kg
- Pantotenat 20 ng/kg
- Vitamin B12 30 g/kg
- Biotin 80-200 g/kg
- Piridoksin 5 mg/kg
- Inositol 10-1000 mg/kg
- Kolin 20 g/kg
Mouse handling and
manual restraint. Apply
slight, rearward traction
on the tail (A). Grasp skin
behind ears with thumb
and index finger (B).
Transfer the tail from the
preferred hand to
beneath the little finger
of the hand holding the
scruff of the neck (C).
From Current Protocols in Immunology
Placing a mouse on a cage lid and grasping the loose
skin behind the ears by the thumb and forefinger
As soon as the mouses head is restrained, the mouse can be picked up and the tail secured within
your ring finger and little finger
From Current Protocols in Immunology
A. Blood collection from tail vein
B. Blood collection from orbital sinus
C. Blood collection from cardiac
puncture
D. Blood collection from saphenous vein
E. Intraperitoneal injection
F. Subcutaneous injection
G. Oral Feeding
H. Sexing
For collection of small amount of blood (Approximate 0.1 ml )
75% alcohol cotton ball for surface
disinfection
Small plastic bottle with 1/2 cm
diameter holes in
both ends as
mouse restrainer
Scissors
Pipetteman and tips
A vial for blood collection
BLOOD COLLECTION FROM TAIL VEIN
OF MOUSE AND RAT USING
MICROHEMATOCRIT TUBE
From Current Protocols in Immunology
Visualize a
sampling site of
the lateral tail
vein at
approximately
midpoint on the
length of the tail.
BLOOD COLLECTION FROM TAIL VEIN
OF MOUSE AND RAT USING
MICROHEMATOCRIT TUBE
From Current Protocols in Immunology
Collect blood
from the hub
of the needle
with the
microhemato
crit tube.
Push the mouse into the restrainer
Leave the tail of the mouse outside
the cover of the restrainer
Amputate the tip of the mouse tail by scissors
Massage the tail and collect blood by pipetteman
Should apply anesthetic before blood withdraw
A convenience and easy apply method for blood collection in mouse
Collect amount up to 0.5 ml
75% alcohol cotton ball for surface disinfection
Hypnorm for general anesthetic
27 G needle with 1 ml syringe for injection
Glass capillary tube and vial for blood collection
Anesthetize a mouse by intraperitoneal
injection of Hypnorm
BLOOD COLLECTION FROM ORBITAL
SINUS OR PLEXUS OF MOUSE AND RAT
Introduce the
end of the
microhematocrit
tube at the
medial canthus
of the orbit.
From Current Protocols in Immunology
BLOOD COLLECTION FROM ORBITAL
SINUS OR PLEXUS OF MOUSE AND RAT
Slowly, and with
axial rotation,
advance the tip of
the microhematocrit
tube gently towards
the rear of the
socket until blood
flows into the tube.
From Current Protocols in Immunology
BLOOD COLLECTION FROM ORBITAL
SINUS OR PLEXUS OF MOUSE AND RAT
Remove the
microhematocrit
tube from orbit
and dab excess
blood from the site
with a gauze
sponge or swab
moistened in
saline or PBS.
From Current Protocols in Immunology
Use a sharp end glass capillary tube to
penetrate the orbital conjunctiva and rupture
the orbital sinus
Collect blood with a vial
For collect up to 1 ml of blood within a short period of time
Must be performed under general anesthetic
75% alcohol cotton ball for surface disinfection
Hypnorm used as anesthetic
27G needle with 1 ml syringe for injection
24G needle with 3 ml syringe for blood withdraw
Anesthetize a mouse by intraperitoneal
injection of Hypnorm
Disinfect the thorax area with 75%
alcohol cotton ball
Search for the maximum heart
palpitation with your finger
Insert a 24G 1 needle through the thoracic wall at the point of maximum heart palpitation
Withdraw blood slowly by your right hand
This method is used of multiple samples are taken in the course
of a day
It can also be applied on rats, hamsters, gerbils and guinea-
pigs
75% alcohol cotton ball for surface
disinfection
50 ml syringe tube with small holes at
the end as
restrainer
a scalpel and shaver for remove of hair
24 G 1 needle for release of blood
tips and pipetteman for blood collection
Placing a mouse on a cage lid and grasping the loose
skin behind the ears with your thumb and forefinger
Place the mouse in the restainer
Pull out the leg and removed the hair by
a assistant
Hair can also be shaved by using a
small scalpel
Apply vaseline after disinfect the surface area to
reduce clotting and coagulation during blood
collection.
Use a 24 G 1 needle to puncture the vein and release blood from the saphenous vein
Use a Microvette or a pipetteman with tip to collect
blood from the saphenous vein
Approximate 100 microliters can be collected
Flex the foot of the mouse to reduce the flow of
blood back to the puncture site
A cotton ball is applied to the puncture site
to stop further bleeding
A common method of administering drugs to
rodents
75% alcohol cotton ball for surface disinfection
25G 1/2 needle with 1 ml syringe for injection
Place a mouse on a cage lid and grasping the
loose skin behind the ears with your thumb and
forefinger
As soon as the mouses head is restrained, the mouse can be picked up and the tail secured within
your ring finger and little finger
The injection site should be in the lower left quadrant
of the abdomen because vital organs are absent
from this area. Only the tip of the needle should
penetrate the abdominal wall to prevent injection into
the intestine.
The most common method for
immunology studies
75% alcohol cotton ball for surface disinfection
25G 1 needle with 1 ml syringe for injection
Pick up a nude mouse and spin its tail to put it in a faint condition
Grasp the loose skin on the back of the mouse from
ears along the legs and restrain the legs with your
ring finger and little finger
After disinfect the surface area, insert the needle in
the lateral side of the abdominal wall and push
upwards to the armpit of the mouse
A lump of injection substance can be
seen through the skin after injection
From Current Protocols in Immunology
1. Fill syringe with
injectate and remove air
bubbles.
Removal of air bubbles is critical to avoid air embolism.
2. Place mouse in a
restrainer.
INTRAVENOUS
INJECTION OF MOUSE
From Current Protocols in Immunology
3. Warm the tail with a
heat lamp or by
immersing in warm water
to dilate vessels.
4. Swab the tail with 70%
ethanol on a gauze
sponge or swab.
INTRAVENOUS
INJECTION OF MOUSE
From Current Protocols in Immunology
5. Immobilize the tail with
gentle traction.
6. Visualize the lateral tail
vein and insert needle
parallel to the vein 2 to 4
mm into the lumen.
Keep the bevel of the needle facing up.
INTRAVENOUS
INJECTION OF MOUSE
From Current Protocols in Immunology
7. Inject slowly.
No bleb should form if
needle was properly
located. If a bleb appears,
indicating failure to
cannulate the vein,
additional attempts may
be made proximally. Thus
it is helpful to make the
first attempt at injection as
close to the tip of the tail
as possible.
INTRAVENOUS
INJECTION OF MOUSE
From Current Protocols in Immunology
8. Withdraw the
needle and apply
digital pressure
if necessary to
achieve
hemostasis.
INTRAVENOUS INJECTION
OF MOUSE
Gastric intubation ensures that all the material was
administered
Feeding amount limited to 1% of body weight
A 18 G stainless steel, ball tipped needle
a glove
Grasp the loose skin on the back of the
mouse and restrain its tail with your ring finger and little finger. Then, introduce the
feeding tube from the pharynx in to the
esophagus when the mouse is in the act of
swallowing.
Common complications
associated with gastric
intubation are damage to the
esophagus and administration of
substance into the trachea.
Careful and gentle passage of
the feeding needle will greatly
reduce these possibilities.
The anatomy picture showed the position of the
feeding needle tip inside the esophagus with the
heart and sternum removed.
Sexing mice - The distance between the anal and genital
orifices is greater in the male (left) compared to the female
(right).
The top two mice are
neonates and
note that the
anogenital distance is
larger in the male than
in the female
neonates, the penis
and vulva cannot be
easily differentiated
and so are referred to
as a genital papilla.
The bottom two animals
are adults; genitalia
are differentiated.
Male Female
Also, nipples become evident in females at about 10 days of age.
Male mice, like other rodent species, retain an open inguinal canal in adulthood. That is, the descended
testicles communicate with the abdominal cavity.
Depending on the position in which the mouse is
held, the testicles may be retracted into the
abdomen or descended into the scrotum.
Because of the open inguinal canal, castration of mice requires that the surgeon use caution when
applying tension to the testicle. Too much tension
can result in the intestines being pulled through the
inguinal canal.
Occasionally people experience difficulty in determining gender in mice at weaning age, although
the anogenital distances are markedly different
between males and females.
Newborns have a subtle difference in the anogenital distance, due to their small size and scale of the
anatomical landmarks. In determining the gender of
newborns, its best to examine several animals side by side to distinguish the males from the females.
The difference becomes more apparent after a few days of age. Another landmark is the presence of
nipples in the females from 10 days of age, which are
absent in the male. Darker mice are more difficult to
differentiate than light colored mice.
Newborn through Day 5
The images on the next slide show pups from
1 to 5 days of age. (The following slides
show mouse pups from 6 to 15 days of
age.)
Newborn, or 1 day old, mice are very red,
helpless, and hairless. Because of their
color and lack of hair, they are often
referred to as pinkies.
Day 6 through 10
The next slide shows images of pups at
ages of 6 through 10 days. The stages
of fur growth are an important
indicator of age during this period of
time
Here are images of pups at ages of 6 through 10 days. The
stages of fur growth are an important indicator of age
during this period of time.
Day 11 through 28
The next slide shows images of pups
from 11 days through 4 weeks of age.
This 18 day period is one of rapid growth
and change as their eyes open and
activity increases.
Carbon Dioxide Asphyxiation
60% to 100% CO2 (tank or house
carbon dioxide) or dry ice Impervious container with raised floor
and lid or CO2 chamber with lid
Carbon Dioxide Asphyxiation
Precharge the impervious container or the
CO2 chamber with CO2 prior to
introducing the animals.
If dry ice is used as the CO2 source, a raised floor is necessary to prevent animal contact with the dry ice. Placing warm water on the dry ice facilitates vaporization and filling of the container.
Carbon Dioxide Asphyxiation
Unconsciousness will occur within 30 sec,
but animals should be left in the container
for several minutes to ensure death.
4. Verify death by lack of cardiac pulse and
fixed and dilated pupils prior to carcass
disposal.
Carbon Dioxide Asphyxiation
Cause perivascular edema
in the lungs or alveolar
hemorrhage. Neonates are
resistant to the effects of
high levels of CO2
Cervical Dislocation of Mouse
Institutional Animal Care
and Use Committee policy.
Cervical Dislocation of Mouse
Remove the mouse from its cage by grasping the base of the tail. Place it
on a smooth, hard surface without
releasing the tail.
Place the pencil or metal rod firmly behind the ears and across the neck.
Pull the tail sharply to the rear while pressing down on the neck (to quickly
dislocate the cervical vertebrae).