Sigma Accustain Trichrome Stain Kit (Catalog #HT15) contains:

Biebrich Scarlet-Acid Fuchsin Solution (# HTIS-1, 0.9% biebrich scarle 0.1% acid fuchsin, 1% acetic acid),
Phosphotungstic Acid Solution (#HTIS-2, 10% phosphotungstic acid),
Phosphomolybdic Acid Solution (#HTIS-3,  10% phosphomolybdic acid ), and
Aniline Blue Solution (#HTIS-4, 2.4% aniline blue, 2% acetic acid)

Weigert's Iron Hematoxylin Set (Sigma catalog #HTI0-79)

Deparaffinize slides and rehydrate sections:
3 x 3'                        ·Xylene (blot excess xylene before going into ethanol)
3 x 3'                         100% ethanol
1x 3'                          95% ethanol
1x 3'                          80% ethanol
1x 5 '                         deionized H20

Stain in Working Weigert's Iron Hematoxylin Solution for 5 minutes.  Make Hematoxylin Solution fresh by adding equal volumes of Solution A (1% Hematoxylin in 95% EtOH) and Solution B (1.2% Ferric Chloride and 1% Acetic Acid in distilled water). The working solution is good for approximately  10 days.
** Hematoxylin stains nuclei blue-black.

Wash in running tap water for 5 minutes.  Rinse in deionized water.

Stain in Biebrich Scarlet-Acid Fuchsin for 5 minutes.
          Decreased red staining usually indicates that the staining solution has aged or been overused and should be discarded.
          ** Beibrich scarlet-acid fuchsin stains cytoplasm and muscle red.

Rinse in deionized/distilled water.

Place the slides in Phosphomolybdic/Phosphotungstic  Acid Solution for 5-10 minutes.  Freshly prepare Working Phosphotungstic/Phosphomolybdic  Acid Solution by mixing 1volume of Phosphotungstic Acid Solution and 1volume of Phosphomolybdic Acid Solution with 2 volumes of distilled water.  Discard after one use.  Formation of a precipitate in Phosphomolybdic Acid Solution does not affect performance.
         ** This allows for uptake of the aniline blue stain.

Stain sections in Aniline Blue Solution for 5 minutes.
         **Aniline blue stains collagen blue.

Rinse slides briefly in distilled water.

Place slides in 1% acetic acid solution for 3-5 minutes.  Discard this solution.
         ** Rinsing in acetic acid after staining renders the shades of color more delicate and transparent.
         ** Ifblue staining of connective tissue appears faded, the section has probably been overdifferentiated in the acetic acid solution.

Dehydrate to xylene.

2 x 3 '         95% ethanol
2 x 3 '         100% ethanol (blot excess ethanol before going into xylene)
3 x 5'          Xylene

Leave slides in xylene overnight to get good clearing of the ethanol.

Coverslip slides using Permount or Polymount (xylene based).

  • Place a drop of Permount on the slide using the glass rod, taking care to leave no bubbles. (don't stir the Permount with the rod too much and make sure that xylenes still cover the slide)
  • Angle the coverslip and let fall gently onto the slide.  Allow the Permount to spread beneath the coverslip, covering all the tissue.
  • Dry overnight in the hood or at 37°C.



      Note: Use only freshly prepared buffers and solutions.

  1. Paraffin removal:
    2 x 5’ in xylene
    2 x 5’ in 100 % EtOH
    2 x 3’ in 95 % EtOH
    1 x 3’ in 70 % EtOH
    3 x 1’ in DD H2O
  2. Antigen Retrieval
    Vector Antigen Unmasking Solution (1:100 in DD H2O, 1 L Vf)
    Prepare 4 bowls with 250 ml each
    Place all 4 bowls in microwave and heat solution in microwave 4 mins at 80% power, keeping solution at 95 oC
    Add slides to first bowl and microwave 4 mins at 80% power, rotate to next bowl and microwave 1 min at 80% power, repeat 3 times at 1 min, 80% power, moving to fresh solution each time.
  3. Allow slides to cool in solution 12 mins
  4. Wash slides 2x2’ in 1XPBS
  5. Optional: Block endogenous antigens and peroxidases with 0.1-1% hydrogen peroxide diluted in PBS for up to 30 minutes
  6. Wash 2 x 4 mins with 1XPBS

    ImmunoCruz ABC staining system:
  7. Incubate sections for one hour in 1.5% blocking serum in 1XPBS.
  8. Blot excess blocking serum from slides.
  9. Incubate sections with primary antibody for 2 hours at room temperature, or overnight at 4 oC. Do not allow sections to dry. Wash with 3 changes of 1XPBS for 5 mins each.
  10. Incubate sections for 1.5 hours with biotinylated secondary antibody. Wash with 3 changes of PBS for 5 minutes each.
  11. Incubate sections for 30 mins with AB enzyme reagent. Wash with thres changes of PBS for 5 mins each.

    Vector NovaRED substrate kit for peroxidase (Cat SK-4800):
  12. Immediately before use on tissues sections, prepare the substrate solution as follows:
              a.  To 5 ml of distilled water add 3 drops of Reagent 1 and mix well.
              b.  Add 2 drops of Reagent 2 and mix well.
              c.  Add 2 drops of Reagent 3 and mix well.
              d.  Add 2 drops of the Hydrogen Peroxide Solution and mix well.
  13. Incubate tissue sections with substrate at room temperature until suitable staining develops (usu 5-15 mins). Longer incubations may increase sensitivity. Wash the sections for 5 mins in tap water.
  14. Counterstain in Gills hematoxylin for 5-10 seconds (may need to filter hematoxylin before use).
  15. Wash 3 x 1 min in tap water.
  16. Dehydrate:
    1 x 5 sec 70% EtOH
    2 x 5 sec 95% EtOH
    2 x 5 sec 100% EtOH
    2 x 5 min 100% xylenes
  17. Apply 2-3 drops permount to section and apply coverslip. Dry overnight.




Fluorescein isothiocyanate–dextran, average mol wt 70,000 (FITC:Glucose = 1:250), sigma Aldrich, 46945. Dissolve the powder with sterile water to make 100 mg/ml stocking solution.

  1. Culture medium change: replace medium with fresh culture medium in inserts and 12 well plate, 500 μl in insert with 1:1000 FITC-dextran (500 μl medium with 0.5 μl FITC-dextran stocking solution) and 1000 μl without FITC in lower chamber. Put back into cell culture incubator and incubate for 24 h.
  2. Fluorescein of medium in lower chamber is read by a SpectraMax M2 Microplate Readers by multipoint with depth check with filters appropriate for 485 nm and 535 nm excitation and emission, respectively
  3. Value of fluorescein is recorded and analyzed.


Equipment: Millicell® ERS-2 Voltohmmeter

  1. Culture medium change: replace medium with fresh culture medium in inserts and 12 well plate, 500 μl in insert and 1000 μl in lower chamber. Put back into cell culture incubator and incubate for 30 min.
  2. Electrode sterilization and neutralization:
    10 min 75% ethanol
    10 min PBS
  3. TER measurement:

TER of each insert is measured by three points (0, 4 and 8 o’clock position). Value of TER is recorded when the number becomes stable on the screen (R1). TER of inserts without cells are considered to be blank control (R2). (S: surficial area of insert membrane)

TER = R2-R1

TERR = (R2-R1)/S

Note: TER: transepithelial resistance; TERR: transepithelial resistance rate; S: surficial area of insert membrane.


RNeasy Kit (Qiagen)
PrimeScript™ RT Reagent Kit (Takara)
SYBR® Advantage® qPCR Premix (Takara)

Total RNA extraction: RNeasy Kit (Qiagen)

1. Place the cell culture plate on ice and wash the cells with ice-cold PBS for 3 times.
2. Aspirate the PBS, then add 350 μl lysis buffer.
3. Pipet the lysate directly into a QIAshredder spin column placed in a 2 ml collection tube, and centrifuge for 2 min at full speed.
4. Add 1 volume of 70% ethanol to the homogenized lysate, and mix well by pipetting. Do not centrifuge.
5. Transfer up to 700 µl of the sample, including any precipitate that may have formed, to an RNeasy spin column placed in a 2 ml collection tube. Close the lid gently, and centrifuge for 15 s at >8000 g (>10,000 rpm). Discard the flow-through.
6. Add 700 µl Buffer RW1 to the RNeasy spin column. Close the lid gently, and centrifuge for 15 s at >8000 g (>10,000 rpm) to wash the spin column membrane. Discard the flow-through.
7. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 15 s at >8000 g (>10,000 rpm) to wash the spin column membrane. Discard the flow-through.
8. Add 500 µl Buffer RPE to the RNeasy spin column. Close the lid gently, and centrifuge for 2 min at >8000 g (>10,000 rpm) to wash the spin column membrane.
9. Place the RNeasy spin column in a new 2 ml collection tube, and discard the old collection tube with the flow-through. Close the lid gently, and centrifuge at full speed for 1 min.
10. Place the RNeasy spin column in a new 1.5 ml collection tube. Add 30–50 µl RNase-free water directly to the spin column membrane. Close the lid gently, and centrifuge for 1 min at >8000 g (>10,000 rpm) to elute the RNA.

cDNA reverse transcription (Takara PrimeScript™ RT Reagent Kit)

11. Reaction mixture preparation:

Reagent Volume Volume Final concentration
5X PrimeScript Buffer (for Real Time) 2 µl 4 µl 1X
total RNA up to 500 ng up to 1000 ng 0.4 µM
RNase Free dH2O      
Total 10 µl 20 µl  

12. Incubate the reaction mixture under the following condition.
37℃ 15 min (Reverse transcription)
85℃ 5 sec (Inactivation of reverse transcriptase with heat treatment)

Real-time PCR (Takara SYBR® Advantage® qPCR Premix)

13. PCR reaction mixture preparation:                                 

 PCR Reaction Mixture
Reagent Volume Volume Final Concentration
SYBR Advantage qPCR Premix (2X) 10 µl 12.5 µl 1X
PCR Forward Primer (10 µM) 0.8 µl 1 µl 0.4 µM
PCR Reverse Primer (10 µM) 0.8 µl 1 µl 0.4 µM
ROX Reference Dye (50X) 0.4 µl 0.5 µl 1X
Template (< 100 µg) 2 µl 2 µl  
dH2O 6 µl 8 µl  
Total 20 µl 25 µl  

14. Incubate the reaction mixture under the following condition

Stage 1: Initial denaturation
Reps: 1
95℃ 30 sec
Stage 2: PCR reaction
Reps: 40
95℃ 5 sec
60℃ 30 - 34 sec
Dissociation Stage
30 sec
Melt Curve Stage



1.  Solutions and reagents:

Lysis buffers:
         NP-40 buffer
‒          150 mM NaCl
‒          1.0% NP-40 (possible to substitute with 0.1% Triton X-100)
‒          50 mM Tris-HCl, pH 8.0
‒          Protease inhibitors

RIPA buffer
‒          150 mM NaCl
‒          1.0% NP-40 or 0.1% Triton X-100
‒          0.5% sodium deoxycholate
‒          0.1% SDS (sodium dodecyl sulphate)
‒          50 mM Tris-HCl, pH 8.0
‒          Protease inhibitors

Tris-HCl: 20 mM Tris-HCl

Protease inhibitors

Running, transfer and blocking buffers
         3X buffer/loading buffer
‒          6% SDS
‒          15% 2-mercaptoethanol
‒          30% glycerol
‒          0.006% bromophenol blue
‒          0.125 M Tris-HCl
Check the pH and adjust to 6.8

         Running buffer (Tris-Glycine/SDS)
‒          25 mM Tris base
‒          190 mM glycine
‒          0.1% SDS
Check the pH and adjust to 8.3

         Transfer buffer (wet)
‒          25 mM Tris base
‒          190 mM glycine
‒          20% methanol
‒          Check the pH and adjust to 8.3

For proteins larger than 80 kDa, we recommend that SDS is included at a final concentration of 0.1%.
         Blocking buffer
3–5% milk or BSA (bovine serum albumin). Add to TBST buffer. Mix well and filter.

2. Preparation of lysate from cell culture
1. Place the cell culture dish on ice and wash the cells with ice-cold PBS for 3 times.
2.  Aspirate the PBS, then add ice-cold lysis buffer (1 mL per 107 cells/100 mm dish/150 cm2 flask; 0.5 mL per 5x106 cells/60 mm dish/75 cm2 flask)
3. Scrape adherent cells off the dish using a cold plastic cell scraper, then gently transfer the cell suspension into a pre-cooled microcentrifuge tube and keep on ice for 10 min.
4. Centrifuge in a microcentrifuge at 4°C, 15,000 rpm for 15 min.
5. Gently remove the tubes from the centrifuge and place on ice, aspirate the supernatant and place in a fresh tube kept on ice, and discard the pellet.
6. Remove a small volume of lysate to perform a protein quantification assay.
7. Boil each cell lysate in 3X sample buffer at 100°C for 5 min. Lysates can be aliquoted and stored at -20°C for future use.

Loading and running the gel
8. Load equal amounts of protein into the wells of the 8-20% SDS-PAGE gel, along with molecular weight marker. Load 20–30 μg of total protein from cell lysate or tissue homogenate, or 10–100 ng of purified protein.
9. Run the gel for 0.5 h at 90 V and then 1 h at 120 V.
The gel percentage required is dependent on the size of your protein of interest:
Protein size     Gel percentage
4–40 kDa        20%
12–45 kDa      15%
10–70 kDa      12.5%
15–100 kDa    10%
25–100 kDa    8%
Gradient gels can also be used.

Transferring the protein from the gel to the membrane

10. The membrane can be either nitrocellulose or PVDF. Activate PVDF with methanol for 1 min and rinse with transfer buffer before preparing the stack. The time and voltage of transfer may require some optimization. Transfer of proteins to the membrane can be checked using Ponceau S staining before the blocking step.

Antibody staining

11. Block the membrane for 1 h at room temperature or overnight at 4°C using blocking buffer.
12. Incubate the membrane with appropriate dilutions of primary antibody in blocking buffer for 2 h at room temperature or overnight at 4°C
13. Wash the membrane in three washes of TBST, 5 min each.
14. Incubate the membrane with the recommended dilution of conjugated secondary antibody in blocking buffer or TBST at room temperature for 1 h.
15. Wash the membrane in three washes of TBST, 5 min each.
16. For signal development, follow the kit manufacturer’s recommendations. Remove excess reagent and cover the membrane in transparent plastic wrap. Acquire image using darkroom development techniques or ChemiDoc for chemiluminescence.



Gloves must be worn throughout the procedure. All work should be done in a fume hood.

  1. Rinse cell monolayers briefly in PBS.
  2. Fix in situ with 2.5% glutaraldehyde in PBS, pH 7.4, for 1 hour at room temperature.
  3. Wash 3 times in PBS buffer (10 minutes each).
  4. Post-fix monolayers for 1 hour at 4° C in 1% OsO4 with 1% potassium ferricyanide.
  5. Wash 3 times in PBS buffer (10 minutes each).
  6. Dehydrate monolayers in a graded series of alcohol (30%, 50%, 70%, and 90% - 10 minutes) with three changes in 100% ethanol (15 minutes each).
  7. Change three times in epon (1 hour each).
  8. Remove last change of epon and invert beam capsules full of resin over relevant areas of the monolayers and polymerize at 37° C overnight and then 48 hours at 60° C.
  9. Pop off the beam capsules and underlying cells from the bottom of the petri dish and section.


Primary BPH stromal cell culture

1. Primary prostate tissue specimens obtained from surgeries are stored in RPMI 1640 medium at 4 °C (recipe below).

2. Wash prostate tissue specimens with 3 ml ice cold PBS 3 times.

3. Mince tissues (into ~2 mm pieces) in a sterile 6 cm dish using sterile scissors. Transfer up to 10 pieces using sterile tweezers into a 1.5 ml Eppendorf tube.

4. Minced tissue specimens (1-2 mg) are digested by 1 hr incubation at 37 °C in 1 ml of 2.4 U/ml Dispase II (Cat # 04942078001, Roche Applied Science, Indianapolis, IN) in RPMI-1640 media on a rocker.

5. Centrifuge minced tissue suspension at 180 x g for 5 mins.

6. Aspirate the supernatant and resuspend cell pellet in 1 ml of 50/50 DMEM/F12 (recipe below).

7. Add 4 ml 50/50 DMEM/F12 to a 6 cm culture dish. Add 1 ml of cell suspension with gentle mixing via repeated pipetting. Total volume is 5 ml.

8. Incubate plated cells at 37 °C, 5% CO2 for 5 days without disturbing. Add 2 ml of fresh medium on top of medium to avoid evaporation on the 5th day. Stromal cell proliferation should be evident by day 7.

9. When stromal cells reach 95% confluence, cells are serially passaged using trypsin:EDTA (0.25%:0.53 mM) solution, then neutralized with 50/50 DMEM/F12. Spin down cells and remove the supernatant from the pellet. Cells are then resuspended with 50/50 DMEM/F12 and split 1:3. Aliquots can be stored at -80 °C as passage 1 (freezing medium is 10% DMSO in 50/50 DMEM/F12) or reserved for subsequent experiments.

10. Primary cells are used in experiments from passage 1-15.

Stromal conditioned medium

1. Seed primary stromal cells (200,000 cells) into 6 cm culture dishes with 5 ml of 50/50 DMEM/F12 and culture to 100% confluence (~ 2 days).

2. Collect conditioned medium (CM) by aspirating from the culture dish into a 15 ml falcon tube and replace with fresh medium (5 ml).

3. Centrifuge the stromal CM at 180 x g for 5 min to remove cellular debris and collect the supernatant. Store the CM at -20 °C.

3D Matrigel culture of prostatic epithelial cells

1. Seed prostate epithelial cells (300,000 cells) (i.e., BPH-1 cultured in RPMI 1640 or BHPrE1 cultured in 50/50 DMEM/F12) into 6 cm plates until 80% confluent.

2. Warm a sterile 24-well plate at 37 °C for 15 mins.

3. Trypsinize epithelial cells with trypsin:EDTA (0.25%:0.53 mM) and neutralize with 50/50 DMEM/F12. Spin down cells and remove the supernatant from the cell pellet. Add 5 ml of DMEM/F12 and determine the number of cells. Based on the total number of cells to be used in 3D culture (i.e., 2,520 cells/well), transfer the proper volume of the cell suspension to a sterile tube. Spin down the cells and remove the supernatant from the pellet. Resuspend cells in Matrigel at a concentration of 63 cells/1 μl of Matrigel. Avoid generating bubbles in the Matrigel.

4. Carefully seed 40 μl of cell:Matrigel suspension in the center of each well in the 24-well plate.

5. Cover the plate and place it upside down in a 37 °C incubator for 15 mins to allow cell:Matrigel suspension to solidify.

6. Carefully add 500 µl stromal CM (see above) to each well. Add the CM along the edge of the well wall to avoid disturbing the Matrigel. Then, add an additional 500 µl fresh medium to each well. Culture the epithelial cells without disturbing for 2 days at 37 °C.

7. Remove media from plate and replace with a fresh mixture of 500 μl CM and 500 μl medium every 2 days.

RPMI 1640 medium
RPMI 1640 medium
1% L- Glutamine
1%  Penicillin-Streptomycin
10% FBS


50/50 DMEM/F12
50/50 Dulbecco's modified Eagles medium (DMEM)/F12
1 µg/ml insulin-transferrin-selenium-X
0.4% bovine pituitary extract
3 ng/ml epidermal growth factor
1% L- Glutamine
1%  Penicillin-Streptomycin
5%  FBS