HistologyHistology kdorfman Wed, 02/01/2012 - 20:56
Histology SuppliesHistology Supplies kdorfman Fri, 05/30/2014 - 15:07
22 x 50 mm 1.5
Krackeler 119-1415-15 $19/10 1oz packs
0.6 mL multicolor microfuge tubes (for aliquoted antibodies)
Krackeler 383-MCT060A Snaplock Microtubes Assorted $61.43 1000/PK
Fisher 05-408-128 Pack of 500 for $35.86 Tube, Microcentrifuge;
Fisherbrand; PP; w/Flat-top snap caps w/insertion spot, Assorted
TBS, inc. Shur/Sharp DMB-LP Fisher 22-047-093
Fisher Scientific* HistoPrep* Stainless-Steel Base Molds (30 L x 24 W x 5mm H) Cat No.: 15-182-505D
MX35 Priemier + ThermoScientific 34degrees, 80 mm
Cellusolve: Ethylene Glycol Monobutyl Ether (Fischer) Cat No.: E179-4
Toluene Fisher T326F-1GAL
PEN CREATE AQUEOUS BARRIER
Quantity: 2 Fisher Cat No.: 23-769-300
Andwin Scientific Tissue-Tek* CRYO-OCT Compound Quantity: 1 case Fisher Cat No.: 14-373-65
Bouin's Fixative Solution (Ricca Chemical) Quantity: 1 Fisher Cat No.: 112016
MOLECULAR BIOLOGY GRADE ETHANOL Quantity: 1 Fisher Cat No.: BP28184
2-BUTOXYETHANOL 99% 1 LITER Quantity: 1 fisher E179-4 Cat No.: AC154330010
mounting medium (SouthernBiotech Dapi Fluoromount-G, Fisher OB010020)
Lab ProjectLab Project kdorfman Fri, 05/30/2014 - 14:11
LAB PROJECT: IMMUNOHISTOCHEMISTRY (original from Bryan)
Objective: To use immunohistochemistry to identify the location of specific molecules and/or structures in organ sections.
1) Every group will use the naturally fluorescent probe, DAPI to stain cell nuclei. In addition, your group should select 2 antibodies or probes to stain the frozen sections of your organ. Be sure to consider the possible secondary antibodies and their fluorochromes when making your selection; one should be marked with a red fluorochrome (rhodamine) and the other with a green fluorochrome (FITC). Predict what the markers will stain in your particular sections. (ex: anti-laminin will stain basal lamina, etc.) Review your staining plan with a TA before proceeding.
2) Get 2 slides of your frozen sections. Label the slides with your group number. Sections should be on the frosted side of the slide with your group number. Flip the slide over so that the reverse side is facing you. On this side of the slide, use a marker to circle the sections. Doing this will enable you to see where your sections are located in order to stain them
3) Fix slides in the fixative, 1% Formalin for 5 minutes. You must prevent your slides from drying from this point on.
4) Rinse slides in Phosphate-buffered saline (PBS) for 5 minutes on a stir plate in an inverted coplin jar with a magnetic stirrer.
5) Dry the edges of the slides outside the hydrophobic barrier with a Kimwipe, taking care not to touch your sections. Use the Pap Pen to make a hydrophobic barrier on either side of your sections (see Figure above). Place these slides in the humidified staining box.
6) On these slides, use a pipet to place enough blocking solution on the sections to cover them. Let sit for 5 minutes.
7) Drain blocking solution from the slides by tipping them on to some Kimwipes and place 100 ul of your primary staining solution on the sections of each slide. Let sit for 45 minutes. While waiting, stain your other immunohistochemistry slides with toluidine blue (see step 12 below) and/or start making your paraffin sections if you haven’t already.
8) Dip incubated slides in fresh PBS to remove the staining solution, then dump this PBS as well. Rinse the slides for 25 minutes in a new PBS solution, as before.
9) Wipe edges of slides carefully and return them to the staining box. Place 100 ul of your secondary staining solution on the sections of each slide. Let sit for 45 minutes in a humidified box.
10) Again, dip slides in fresh PBST to remove the staining solution, then dump this PBS as well. Rinse the slides for 25 minutes in a new PBS solution, as before.
11) Wipe edges of slides and place one drop of mounting solution on sections. The mounting solution contains DAPI, the cell nuclei marker. Place coverslip on slide. Fix in place with nail polish.
12) Toluidine Blue staining: Fix and rinse the other slides following steps 3-5 above. Place a drop of toluidine blue solution on the sections. After ~15 seconds, rinse the slide and look at it under low power using your microscope. If it is not well stained, stain again. If it is over stained (too dark) use another slide and start staining again for a shorter time. Consult with Bryan.
13) Image your fluorescent and toluidine blue-stained slides.
|Antibody Name||Antigen recognized||Produced in||Dilution|
|DHB 2E8 (IgG2a)||Laminin||Mouse||1:2|
|DHB Ralph 3.1 (IgG2a)||Integrin, alpha-3||Mouse||1:2|
|DHB M3F7 (IgG1)||Collagen Type IV||Mouse||1:2|
|DHB II-II6B3 (IgG1)||Collagen type II||Mouse||1:2|
|DHB JLA20 (IgM)||Actin||Mouse||1:2|
|DHB SV2||Synaptic Vesicles||Mouse||1:2|
|DHB TI-4||Anti-Troponin I||Mouse||1:2|
|DHB B4-78||Anti-Alkaline Phosphatase||Mouse||1:2|
|DHB T14||Anti-Myosin Light Chain||Mouse||1:2|
|Antibody Name||Produced in||against||Dilution|
|F0382-1ML FITC CONJUGATE||GOAT||ANTI-RABBIT IGG||1:80|
|T5393-.5ML T5393||Goat||Anti-Mouse IgG||1:64|
Making Composite ImagesMaking Composite Images kdorfman Wed, 04/19/2017 - 17:14
- Open Image J from the Applications Menu in the Finder Window
- Open all 3 of your images by dragging their icons to the ImageJ toolbar.
- Open the Image menu, and select color>merge
- Put your rhodamine (phalloidin) image in the red channel, your fluorescein (FITC) image in the green channel, and your DAPI image in the blue channel. Leave the other channels blank.
- Save the composite image both as a TIFF and a JPEG with a reasonable file name, but don't discard the originals. The TIFF will work better if you decide to do further analysis in ImageJ, but the JPEG may work better for your lab report.
If you took the three images without moving the specimen, this image should show you the relative distributions of dsDNA, actin, and the cellular protein your chosen antibody bound to in the cells.
Paraffin Embedding (Rats)Paraffin Embedding (Rats) kdorfman Fri, 05/30/2014 - 18:24
Do all of the following at room temperature.
1.) Fix in Bouin’s (Bouin's Fixative Solution (Fischer) Cat No.: 112016) overnight in tubes.
This step should be kept short and should not include agitation. I have noticed that the intestine and liver could both benefit from shorter fixation times.
2.) Place tissue in cassettes and rinse for 2 hours in running water, then place into 70% ETOH (they may be stored here for up to several months).
There is a large jar with a lid to perform all of the following steps in.
3.) Cellosolve (Ethylene Glycol Monobutyl Ether (Fischer) Cat No.: E179-4) I – 1-2 hours, or longer.
4.) Cellosolve II – 1-2 hours, or longer.
5.) Cellosolve III – 1-2 hours, or longer.
6.) Cellosolve IV – Overnight.
7.) Toluene I – 1 hour.
8.) Toluene II – 1 hour.
9.) Paraffin I – 1 hour.
10.) Paraffin II – 1 hour.
11.) Paraffin III – 1 hour under vacuum (vacuum not necessary).