Kuliah 2. Mencit Sebagai Hewan Laboratorium_2

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).


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

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


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


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.




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.




Remove the

microhematocrit

tube from orbit

and dab excess

blood from the site

with a gauze

sponge or swab

moistened in

saline or PBS.


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


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


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


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


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


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


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


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


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


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.


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.


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).

Kuliah 2. Mencit Sebagai Hewan Laboratorium_2

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