university of california, san diego histology shared resources

Nissi M. Varki M.D.,
Director. Email:
nvarki@ucsd.edu Phone:
858-534-5611
UNIVERSITY OF CALIFORNIA, SAN DIEGO
HISTOLOGY SHARED RESOURCES
9500 Gilman Drive, Mail Code 0687, La Jolla, CA 92093-0687
Phone: 858-822-1011. FAX: 858-534-5611
Program in Vascular
and Blood
Glycosylation. Contact:
858-534-2945 Lisa
Wiggleton BS EMT.
Email:
LWiggleton@ucsd.edu
Consortium for
Functional Glycomics
Contact: Y.Lucie Kim BS
HTL Email:
ylkim@ucsd.edu
Cancer Center
Histology: Contact:
MaryAnnLawrence HT
Laarni Gapuz MT
Sabrina Schuck
Wess Jordan
Gastroenterology /
Mucosal Immunology
Program: Contact:
858-534-4625
Steve Shenouda BS
sshenouda@ucsd.edu
Figure Legend: The figure depicts the recommended arrangement of tissues harvested
from littermate controls and from genetically altered animals, on slides for microscopic
examination in order to visualize and digitally photomicrograph if histo-pathologic
abnormalities are observed.

Tissues and cell types screened during histopathologic analysis:
I. Brain: gray and white matter, endothelium, choroid plexus, pituitary, ventricles olfactory lobes, corpus
callosum, hippocampus, midbrain, cerebellum, choroid plexus, pia mater
II. Heart and circulatory system: cardiomyocytes endothelium and vessels, pericardium of right ventricle,
interventricular septum, cardiac muscle, valves, endothelium.
III. Lungs: bronchiolar epithelium, vessels and endothelium, stroma, mesothelial layer
IV. GastroIntestinal Tract:
A. Liver: hepatocytes, Kupffer cells, bile ducts, portal triads and central veins
B. Pancreas: acini, Islets (endocrine), ducts, blood vessels and lobules
C. Salivary glands, nerves, ganglions, stroma
D. Intestines: all areas: mucosal epithelial cells, smooth muscle layers,
ganglion cells, blood vessels,
a. Esophagus: in addition :
squamous and glandular epithelial mucosal layers
b. Stomach: in addition :mucosal epithelial cell types
c. Duodenum: in addition : Brunners glands
d. Ileum: in addition : villi, goblet cells, Peyer’s Patches
e. Colon: in addition : goblet cells, Peyer’s Patches
V. GenitoUrinaryTract
A. Kidney: Cortex/medulla---Glomeruli, tubules—
proximal/distal, blood vessels
B. Bladder: epithelium, muscle, blood vessels
C. Ovaries/Uterus/Fallopian tubes
D. Placenta
E. Prostate: tubulo-alveolar glands fibromuscular stroma
F. Testis, seminiferous tubules, Sertoli cells and germ cells with mature sperm,
Leydig cells, vessels, epididymis,
VI. Hematopoietic system:
A. Spleen: lymphoid nodules, red pulp, macrophages, megakaryocytes, vessels
B. Thymus: Cortex, medulla, blood vessels
C. Lymph nodes: cortex, medulla, blood vessels, HEVs
D. Bone Marrow
VII. Endocrine: Adrenal, islets in Pancreas, Parathyroid, Thyroid
VIII. Skin and Integument
A. Epidermis, dermis with sweat glands, sebaceous glands and hair follicles
B. Breast: lobules and ducts
C. Bone/ cartilage and fat and skeletal muscle

Tissue Fixation Procedure for Paraffin Processing
The following protocol is to be used for tissue sample processed for light microscopy.
Tissue fixation
1. The tissue removed from the animal should be placed into marked cassettes and them immediately
placed into a container of 10% neutral buffered formalin.
2. Cassettes are to be specially marked with Secureline marker or pencil exclusively as all other marking
can be removed from solvents during processing.
3. The volume of fixative must be 15-20 times over the sample volume and the tissue must be
completely immersed in the solution.
4. Tissue can remain in fixative at ambient temperature up to 48 hours. No agitation is required
5. Check for clarity of formalin solution day to day. If solution is cloudy, change out the old solution
with fresh, especially if tissues will not be processed immediately.
6. The recommended duration of fixation for general mouse tissue that is no larger than 1.5 cm thick is
24- 48 hours. Larger tissue sections should be monitored and fixed for longer periods of time.
7. If glycogen studies are to be made on liver specimens, the fixation time are to be closely monitored or
cytoplasmic vacuolation may be conspicuous.
8. Prior to cassetting or prior to processing, tissue samples are to be trimmed for optimal processing and
embedding. This size restriction is confined to a maximum height of 3mm and tissue must be in the
tissue cassette loosely for ample penetration of paraffin during the processing schedule.
9. Up to one day prior to processing, the tissues can be immersed in 70% alcohol. This step
will halt formalin fixation and acclimatize tissue to the first stage of the dehydrating process.
However, prolonged exposure to 70% alcohol may have a drying effect on small specimens.
10. Spleen and pancreas should be enclosed between cassette sponges, in cassettes, before fixation to
ensure even fixation and sectioning
11. Embryos of differing sizes require specific fixation protocols. Please contact the histology personnel
for help prior to harvesting embryos for microscopic examination
Recommended Materials:
Securline Marker—Fisher Cat. No. 14-905-30
Buffered Formaldefresh—Fisher Cat. No. SF93-4
Freshly prepared 4% paraformaldehyde
Bouin’s solution: Sigma Cat. No. H10-1-32 should be used to fix opened embryos and other small slices
of organs that need good morphologic analysis.
Fix for 6- 8 hours and then transfer to 70% alcohol to “wash out” the yellow before submitting to
histology for automated dehydration, paraffin embedding, sectioning and staining.
ZincFormaldehyde fixative: to fix organs that will be used for immunohistochemistry stains. Fix thin
slices in this fixative obtained from American Master Tech Phone: 209-368-4031. Cat. No. FXZFO

Dissection Protocol
Equipment and Materials:
Styrofoam board or cork cutting board and pins, flat for cutting
Scissors, delicate ½”—Fisher Cat. No. 08-951-5
Bell jar with lid
200ul Pipetman and pipets
Kimwipes, Lab liners, paper towels
Forceps, curved—VWR Cat. No. 25714-007
Secureline Marker—Fisher Cat. No. 14-905-30
1ml syringe—Fisher Cat. No.
Vacutainer 23G ¾ --Fisher Cat. No.0266539
HistoPrep Tissue Capsules—Fisher Cat. No. 15-182-219
Peristaltic Pump
Phosphate Buffered Saline
Isoflurane—MWI Cat. No. 012497
10% Neutral buffered formalin (Buffered Formaldefresh)--Fisher Cat. No. SF93-4
labeled specimen cups
Personal protective equipment eg. lab coat, gloves, eye protection.
Check out this website: www.eulep.org for photographs and other details
Before Autopsy:
1. Determine which organs will be harvested and formalin fixed or frozen.
2. Gather materials and label all necessary cassettes or capsules (see Blocking Chart Protocol.)
3. Record the identifying data on each animal including its appearance and behavior.
4. Record the weight of the animal, the individual organs, any abnormal gross pathology, and the
carcass after the organs are removed. (Relative organ weights can be found on page_____.) If
skeletal defects are suspected save the carcass and obtain a radiograph of the skeleton. The carcass
may be “cleared” using potassium hydroxide and stained with Alizarin Red and Alcian Blue,
following established protocols.
5. Anesthetize and euthanize animals humanely according to animal welfare guidelines.
6. Wet the animals' fur with 70% ethanol to minimize contamination with hair and lay upon a clean
paper towel. Pin extremities to styrofoam or cork board. Dedicated scissors are recommended for
cutting skin and bone.
7. Start from the abdomen and make and a Y-shaped incision should be made to open up the thorax
and abdomen. The ribcage should be opened bilaterally, exposing the underling thymus, heart and
lungs.
8. If the plan is to perfuse the entire animal, canulate the left ventricle of the heart with the catheter
connected to the peristaltic pump and secure the catheter with alligator clips..
9. Make an incision in the right atrium to allow the circulating fluids to exit the right atrium.
10. Pump PBS through the circulatory system, using a peristaltic pump. This will flush out blood.
Adequate perfusion is indicated when the liver and kidneys become pale.
11. Follow the PBS flush with freshly made 4% paraformaldehyde for 15 min.
12. Just before finishing the perfusion, insert a formalin filled syringe into the trachea just above the
lungs until full. Then tie off the trachea to keep the fixative in the alveoli
13. Once perfusion is complete, remove the organs listed below in the order listed. This will ensure
that all the necessary structures are collected.

14. While harvesting the organs, place smaller specimens and the spleen, between sponges in labeled
cassettes before placing them in formalin.
15. The larger structures such as brain, lung, liver, can be placed directly into the fixative. (The brain
needs to be divided into three parts after fixation, so it is recommended that it is placed directly in
fixative, after removal.)
EMBRYOS: Specific fixation methods are recommended for examination of embryos
Please contact personnel in the core lab to plan optimal methods. Really small embryos should be oriented
and embedded in 1% agarose, before being fixed in freshly prepared paraformaldehyde for processing
into paraffin
Older embryos should be dissected open along the abdominal wall so that fixative will enter. Bouins’
fixative is preferred for fixation of older embryos
Any age of embryo may be frozen using the method given here, for frozen sections and
immunohistochemistry.
ADULT ANIMAL necropsy and gross examination:
Remove:
Salivary Glands: located just below the skin under the chin, above the thorax. Lift to take both lobes
whole including attached lymph nodes and place in fixative.
Lymph nodes: The adjacent lymph nodes may be harvested at this time and placed in a cassette along
with the adrenals—the small sized organs process and cut easier this way. Dissect lymph nodes from
behind the elbow, from the mesenteric fat and also from the hilar region of the lungs
Thymus: The thymus has two lobes and sits on the superior and ventral aspect of the heart, just under the
ribcage. It involutes in older animals and becomes increasingly surrounded by fat (older than 3 months).
Be sure to detach and place between sponges in a cassette before fixing so it will fix without curling
around the edges.
Spleen: Is found on the left lateral aspect of the body, below the ribcage, as an elongated dark red-brown
structure. Be sure to detach and place between sponges in a cassette before fixing so it will fix without
curling around the edges
Lungs: The trachea is exposed and the fixative is infused so that both sides of the lungs inflate well. After
fixation, the lobes of the lungs are dissected away from each other and laid out in labeled cassettes so that
sections through each lobe can be made for histopathologic examination.
The trachea, hilar lymph nodes, thyroid and parathyroid may be left to be fixed to be submitted in
additional cassettes
Heart: The distal half of the heart is to be sliced and that portion is to be embedded flat, in order to be
able to compare the thickness of the heart muscle from each ventricle.
Adrenals: The adrenals should be on the superior pole of each kidney and look like little pyramids of
pinkish fat. Detach the each kidney with the adrenals intact and place in a cassette. (The adrenals may
also be removed but care must be taken to place them between two sponges in a tissue cassette before
placing in fixative.)
Pancreas: Is attached to the spleen and may be harvested when removing the spleen.

Liver: Take a large lobe of the liver with gall bladder and slice it flat and fix whole
Ovaries, Uterus, Vagina, Bladder: The uterus can be identified from the two horns which extend up from
the cervix into the pelvis and ends with the ovaries that are just below each kidney. Cut across the cervix
and lift out the uterus with forceps.
Testes: Testes should be located and taken out whole as gently as possible so as to prevent them from
rupturing. The epididymis may be taken along with them especially if the study requires their analysis.
Prostate: The prostate is just underneath the bladder in the males. It can be taken whole with the bladder
to be fixed together for easy identification at the time of trimming for tissue processing.
Intestinal Tract (Stomach, Ileum, Cecum, Colon, Rectum): Grasp the abdominal end of the esophagus
and begin to disembowel the GI tract until the rectum is reached. Care should be taken not to stretch out
the mesenteric fat so that harvesting of the mesenteric lymph nodes can be possible after fixation.
The small intestine should be sectioned into 4-5 parts before immersion fixation.
The COLON is dissected away from the rest of the bowel and opened along the anti-mesenteric aspect.
Taking a swab stick, roll up the FRESHLY opened colon onto the stick so that a compact roll is the end
result.
DO NOT TRY TO ROLL after rinsing the gut in PBS, it will not stick together to make a good roll.
Always use the same method to roll, anal end first always OR small intestine end first, on the inside of the
roll.
Fix the rolled up colon on the stick, in a 15 ml tube with fixative and submit for histologic embedding
and sectioning.
rolled up colon
on a stick
to be immersed
in fixative
Mammary tissue: The inguinal and axillary mammary glands are visible as yellow fat pads adherent to the
underside of the skin beneath a fascial plane. They should be dissected away from the overlying skin.
Skeletal muscle: A small piece of the abdominal wall may be sectioned. If fixing entire carcass, it can be
left intact and a piece can be cut during trimming.
Bone: Turn the mouse over onto its ventral surface. Using forceps and scissors (scalpel is also acceptable)
lift the skin off the left hind and snip off. This will make the bicep femoris and adductor muscles more
readily visible. Cut the limb off right at the juncture of the hip socket and ankle. Repeat this procedure
with the right limb.
Skin: A snip of one of the ears at least half way down to the juncture of the scalp will be sufficient.
Brain: FOR OPTIMAL MORPHOLOGY< it is best to perfuse the animal with freshly prepared 4%
paraformaldehyde before removing the brain for further fixation, before processing into paraffin blocks.
Grasp scalp on the crown of the head with forceps and snip off skin to see skull clearly (see page 9 and 10
of Atlas). While grasping the orbital of each eye with forceps, move scissors slightly underneath the skull
, and run tip of scissors between skull and brain, keeping scissors horizontal, along the plane of the cutting

oard, on both sides of the skull. This will loosen and separate the calvarium from the brain, which can
now be lifted off. Gently lift the brain off the base of the skull with blunt scalpel, carefully making sure
not to rupture the integrity of the brain. (Remove in one piece the cerebrum, cerebellum and as much of
the hind brain as possible.) The pituitary may also be removed for examination at this time. Eyes may be
examined and other parts if necessary.
If desired, the rest of the mouse carcass can now be immersed whole into the formalin solution to retrieve
finer structures during trimming e.g. lymph nodes.
If bone and cartilage are to be examined further, it may be necessary to treat the adult carcass, after
removal of skin, with potassium hydroxide and stain with Alizarin Red/ Alcian Blue, following
established protocols. 2
1. 1 A Color Atlas of Sectional Anatomy of the Mouse, Takamasa Iwaki, Hiroshi Yamashita,
Toshiyuki Hayakawa
2. Manipulating the Mouse Embryo. Hogan, Beddington, Constantini, Lacy. Cold Spring
Harbor Laboratory Press
Others
a) Pathology of the mouse. Ed. Robert R. Maronpot. Cache River Press
ISBN: 1-889899-02X
b) di Fiore’s Atlas of Histology. Victor P. Eroschenko Lee and Febiger
c) Histology. A text and Atlas. Ross, Romrell and Kaye. Williams and Wilkins
ISBN: 0-683-07369-9
d) Human Histology. Stevens and Lowe. Mosby. ISBN: 0-7234-2485-3
e) An Atlas of Histology. Zhang. Springer. ISBN: 0-387-94954-2
f) Pathologic Basis of disease. Robbins,Kumar and Collins. Saunders.
ISBN: 0-7216-7335-X
g) The Laboratory Mouse. Suckow, Danneman, Brayton. CRC Press
ISBN: 0-8493-0322-2
h) A Color Atlas of Sectional Anatomy of the mouse. Iwaki, Yamashita, Hayakawa.
Braintree Scientific Inc. ISBN: 4-900659-58-4
i) The Atlas of Mouse development. Kaufman. Academic Press.
ISBN 0-12-402035-6
j) The House Mouse. Atlas of Embryonic development. Theiler. Springer-Verlag
ISBN: 0-387-05940-7
k) Manipulating the mouse embryo, a laboratory manual. Hogan, Beddington, Costantini,
Lacy. Cold Spring Harbor Laboratory Press ISBN: 0-87969-384-3

Decalcification Protocol
The following protocol is to be used for bone sample processed for paraffin embedded light microscopy
with H&E staining.
Equipment and Materials:
Styrofoam board or cork cutting board
Scissors, delicate ½”—Fisher Cat. No. 08-951-5,
Or Razor Blades, or scalpel
Forceps, curved—VWR Cat. No. 25714-007
Secureline Marker—Fisher Cat. No. 14-905-30 (pencil is also acceptable)
HistoPrep Tissue Capsules—Fisher Cat. No. 15-182-219
Cal-Ex –Fisher Cat. No. CS510-1D in glass jar or specimen cup
Or Cal-Ex II –Fisher Cat. No. CS511-1D in glass jar or specimen cup
Personal protective equipment e.g. lab coat, gloves, eye protection.
Bone decalcification
Generally the hind limb is taken during autopsy and fixed in formalin for a minimum of two days. At the
time the organs are being trimmed for processing, it will be useful to prepare the bone for decalcification
at this step.
1. With a scalpel or other cutting instrument, remove all of the skin and muscles taking care not to
remove the cartilage around the knee joint or scraping the bone too hard. (The skeletal muscles must
be removed as the decalcifying solution does not penetrate through the muscle.)
2. Place the bone in the appropriate cassette (see Blocking Chart) and immerse in container of Cal-Ex at
room temperature. No agitation is required but can be useful.
3. If the limb is not prefixed, it may be placed directly into Cal-Ex II. This solution is a combination of
fixative and decalcifyer and takes no more that two days for both processes to take place. As above,
the skeletal muscles and skin must be removed for the solution to take effect and placed in the
appropriate cassette.
4. For mouse bones, decalcification should not go beyond 2 days as overdecalcification will occur, tissue
basophilia will be lost, and poor hematoxylin staining will result.
5. To halt the decalcification process and to prepare specimen for processing, wash in running tap water
for approximately 2 hours.
6. The tissue is now ready to be placed in 70% alcohol and ready for processing.
As with the other mouse organs, it recommended that the bones do not remain in alcohol longer that
overnight.

MOUSE ORGAN HARVEST and FREEZING
TO FREEZE TISSUES FOR FROZEN SECTIONS:
1. Get dry ice
2. Get plastic bag with tag to dispose off animal when done
3. Get instruments ready and small plastic jar with water to rinse them out in
4. Label plastic bags to be used to contain frozen blocks
5. Lay out vinyl molds as needed with label
6. Make 1:1 OCT: PBS for tracheal infiltration to inflate lungs before freezing
7. Get bench area ready with absorbent material, styrofoam board and needles to pin down
8.
9. In a small styrofoam flat container, make a slurry with small bits of dry ice and 2 methyl
butane. This will be the freezing mixture which will freeze the organs in OCT for frozen
sections
10. Lay euthanized animal on dissecting board
11. Use an inverted Y-shaped incision to open under the skin
12. Harvest lymph nodes from cervical area and elbow and inguinal , pat dry on paper towel
13. Expose trachea and infiltrate lungs with OCT/PBS mixture. Remove entire block and place on paper
towel to drain before freezing flat in vinyl mold with OCT
14. Harvest tissues and place in sequence onto paper towels to dry off tissue fluids before placing into
vinyl molds, flat on the bottom and then fill with OCT. This keeps the tissues all at the same level:
A. spleen/thymus, lymph nodes. Adrenal
B. Liver, pancreas, kidney
C. Heart, OCT inflated and drained lungs
D. GI (small intestine, colon, stomach) and GU (Bladder, prostate, testis, ovary, uterus)
E. Skin, skeletal muscle, salivary glands, mammary glands, (decalcified femur for blocks)
F. Brain
IT IS BETTER TO KEEP THESE AS SHOWN. CAN COMBINE A,B,C and D,E,F WHEN
CUTTING FROZEN SECTIONS SO THAT YOU WILL GET 2 SLIDES PER ANIMAL
15. Place plastic molds containing organs and OCT into dry ice/ isopentane slurry till OCT turns white
16. Remove plastic molds with frozen organs from dry ice/ 2 methyl butane and let them sit for a minute
or so on paper towels, snap off the plastic molds and place frozen blocks into labeled bags for storage
in boxes in the second container of dry ice. All boxes will be stored at minus 70.
SUPPLIES:
1. OCT Compound: VWR Cat No: 25608-930
2. Vinyl molds: Fisher Cat. No 15-182-501D
3. Frozen sample write-on bags: Fisher Cat. No: 01-812-6B
4. 2 methyl butane: Fisher Cat. No. 03551-4

Hematoxylin and Eosin staining of Frozen sections:
1. Fix frozen sections in 10% buffered formalin (Fisher SF93-4) for 20 minutes and wash in water
2. stain nuclei by immersing in GILL II HEMATOXYLIN (Surgipath 01522) for 3 minutes.
3. wash in water and immerse in blueing agent (Fisher CS410-4) for 30 seconds, and wash in water
4. Immerse in 95% ethanol briefly
5. Stain cytoplasm by immersing in EOSIN (Surgipath 01602) for 3 minutes and dip once quickly in
water
6. dehydrate by immersing in 1 changes of 95% ethanol for 1 min, and then in 3 changes of 100%
ethanol (Fisher A962p) for 10 dips each.
7. Immerse in 3 changes of Citrosol (Fisher 22-143-975) for 2 minutes each
8. Mount slides with Cytoseal 60 (VWR 48212-154) using glass coverslips (Surgipath 00145)
Hematoxylin / Eosin staining of PARAFFIN sections:
1. Deparaffinize by immersing successively, in 2 changes of xylene (Fisher X3P) for 10 minutes each
2. Rehydrate by immersing in decreasing concentrations of ethanol (Fisher A962p) :
100% for 5 minutes
100% for 5 minutes
95% for 5 minutes
95% for 5 minutes
3. Immerse in water for 5 minutes
4. Follow steps 2 through 8 above.
1. Hematoxylin and eosin stain on frozen sections for Laser Capture microscopy
(American Journal of Pathology February 2000 page 446):
1. Thaw frozen sections ( on plain untreated glass slides) one at a time to decrease degradation
2. Fix with 70% ethanol for 10 seconds
3. wash in deionized water
4. immerse in fresh MAYERS hematoxylin for 30 seconds
5. wash in water
6. Immerse in blueing solution for 30 seconds
7. wash in 70% ethanol
8. immerse in eosin for 90 seconds
9. dehydrate sections with two 10 second washes in 95% ethanol and two 10 second washes with
100% ethanol
10. place in xylene for 30 seconds

Date:
Immunohistochemistry checks
Aim:
Positive control:
Negative control:
Positive reagent control:
Negative reagent control:
Tissue sections: Frozen Paraffin
Fixation:
Deparaffinization
A. Acetone 10 minutes
Immediate PBS washes Antigen retrieval: Yes No
Cover sections with 1% BSA/PBS
B. 4% fresh paraformaldehyde 0.1%trypsin or trypsinEDTA
heat in O.1M citrate, cool 20 minutes
0.01% TritonX100, PBS washes
---Blocking:
-endogenous peroxidase: 0.03% 0.3% H 2 O 2 in PBS for 20 minutes,
followed by PBS washes
-endogenous biotin: 0.1% avidin 10 minutes, wash;
0.01% biotin 10 minutes, wash
---Block endogenous collagen binding: 1% BSA/PBS;--- add 10% serum
---Primary reagents: Dilutions time incubation
1._____________________________________________________
2_____________________________________________________
3._____________________________________________________
4. ____________________________________________________
5.._____________________________________________________
6. ____________________________________________________
-----Wash in buffer x3
---Secondary reagents Dilutions time incubation
Catalog number and lot number
-----Wash in buffer x3
---Tertiary reagents: Dilutions time incubation
Catalog number and lot number
-----Wash in buffer x3
-----Further tiers and enhancements: Describe
------Chromagen or Fluorescence marker used :
------Wash in buffer x3
------Mayers hematoxylin or nuclear fast red or none:
-----Wash in buffer x3
------Aquamount with or without diluted DAPI