Isolation of Primary Murine Skeletal Muscle Microvascular Endothelial Cells

All animal experiments were approved by the local authorities and conducted according to the German animal welfare act (84-02.05.20.13.097).

1. General Remarks on Animal Experiments

1. Perform all mouse experiments in accordance with the guidelines of the respective institutional animal care and use committee.
2. Keep mice under standardized conditions and according to international guidelines such as the Federation for Laboratory Animal Science Associations (FELASA).
   NOTE: In general this isolation technique can be used for mice independent of age, gender or genetic background. To obtain a sufficient cell number, 4–10 week old males are preferred, because biological properties can vary with age and gender.

2. Preparation of Solutions, Media and Coating

1. Prepare digestion solution (DS) by mixing 2.2 mL of Dulbecco´s Modified Eagle Medium (DMEM) with 200 μL Collagenase-Dispase and 45 μL Desoxyribonuclease (DNase).
2. Prepare 500 mL of Endothelial cell medium (ECM) by mixing 450 mL of DMEM with 50 mL FCS (approximately 10%), 0.25 mL basic fibroblast growth factor bFGF (20 μg/mL; approximately 0.05%) and 5 mL penicillin-streptomycin (approximately 1%). Perform sterile filtration in a glass tube (diameter of filter pores: 0.2 μm). Afterwards store the solution at 4 °C.
3. Coat cell culture plates as described below.
   1. For cultivation of freshly isolated primary murine MEC cover the whole surface with 1 mL per well of a 6 well cell culture plate with a gelatin based speed coating solution (see Table of Materials) according to manufacturer’s instructions for 3–5 min at room temperature (RT).
   2. Aspirate and discard coating solution. Rinse each well with 2 mL sterile phosphate buffered saline (PBS), pH 7.1–7.5 in two consecutive washing steps.
   3. Aspirate and discard PBS. Fill up wells with 1 mL volume of ECM.

3. Isolation of Primary Murine Muscle Microvascular Endothelial Cells (MMEC)

1. Euthanize one adult 4–12 weeks old, male mouse by cervical dislocation without any anaesthesia. The aim is to quickly separate the spinal cord from the brain so as to provide the animal with a fast and painless death.
2. Sever the extremities.
   1. Use a surgical sharp/blunt (cutting edge 42 mm, straight) and sharp/sharp (cutting edge 23 mm, straight) scissor, a straight (serrated, 2 mm x 1.25 mm x 12 cm) and a curved (serrated, 1.3 mm x 1 mm x 13 cm) forceps.
   2. Disinfect all surgical instruments with 70% ethanol.
   3. Position the animal on its back, moisten the legs with 70% ethanol. Sever the whole leg by cutting at the hip joint with the sharp/blunt surgical scissor. Place the extremities in a closed cell culture dish (35 mm x 10 mm).
      NOTE: From now on perform every step under a sterile laminar flow hood and work with sterilized instruments.
3. Isolate muscle tissue (preferentially use the musculus quadriceps femoris or musculus triceps surae from both legs).
   1. Cut the skin open from the hip to the toe-tip by using a sharp/sharp scissor and curved forceps. Hold the toe or footpad with the straight forceps. Peel off the skin with the curved forceps from the toe to the hip.
   2. Isolate the musculus quadriceps femoris by cutting the tendon from the knee and sever the muscle along the femur to the hip. Isolate the musculus triceps surae by severing the Achilles tendon. Hereafter, cut along the tibia to the popliteal fossa and remove the muscle.
      NOTE: If large vessels (A. femoralis, A. tibialis anterior, A. tibialis posterior, A. fibularis) are visible in the isolated muscles, remove vessels to avoid contamination by macrovascular endothelium.
   3. To remove large vessels, hold the part of muscle not containing large vessels with a curved forceps and cut it off next to the vessel. For this protocol, 1 g or less muscle tissue equal to both quadriceps femoris and triceps surae is recommended.
   4. Add 2,445 µL DS (from step 2.1) to a cell culture dish (35 mm x 10 mm) and determine the weight.
   5. Transfer all muscle pieces to this cell culture dish containing the 2445 µL DS and determine the weight. The difference of both measured values provides the dry weight of muscle tissue which must not exceed 1 g.
   6. Cut the whole muscle tissue into small pieces (≤2 mm cubes, about 100 pieces) by using the sharp/sharp scissor.
4. Dissociate the muscle tissue.
   NOTE: This step takes about 120 min.
   1. Store muscle/DS suspension at 37 °C (incubator with 5% CO₂) for 1.5 h. Mix the suspension carefully every 20 min for about 5 min using a 1 mL insulin syringe.
   2. Transfer suspension to a 70 μm nylon cell strainer placed on a 50 mL tube and collect the flow through. Wash the cell strainer with 8 mL of DMEM and collect the flow through.
   3. Discard cell strainer and centrifuge suspension for 10 min at 300 x g at 20 °C. Carefully remove supernatant.
      CAUTION: pellet is easy to lose.
   4. Resuspend cell pellet in 1 mL of an Ammonium-Chloride-Potassium (ACK) lysing buffer (see enclosed Table of Materials) for the lysis of red blood cells and incubate for 30 s at RT. Add 9 mL DMEM + 10% FCS to stop the reaction and transfer cell suspension to a 15 mL tube.
5. Deplete CD45⁺ cells following the CD45 microbeads manufacturer’s instructions. For all following steps of CD45⁺ depletion use MCS buffer (see enclosed Table of Materials).
   NOTE: This step takes about 60 min.
   1. Determine cell numbers using a Neubauer cell counting chamber (expect about 5 x 10⁶ cells per g muscle tissue).
   2. Centrifuge suspension at 300 x g for 10 min at 4 °C. Take off supernatant completely and resuspend cell pellet in 90 µL MCS buffer (per 10⁷ cells or less). Add 10 μL of CD45 microbeads (per 10⁷ cells or less).
   3. Mix cell suspension and incubate for 15 min in the fridge at 4–8 °C.
   4. Add 1mL MCS buffer (per 10⁷ cells or less). Centrifuge at 300 x g for 10 min at 4°C. Remove the supernatant completely and resuspend cells in 500 μL MCS buffer.
   5. For magnetic cell separation use a large magnetic column (LMC) with a reservoir volume of 8 mL and a capacity of up to 2 x 10⁹ total cells. Position the LMC in the magnetic field of the separator.
   6. Rinse with 3 mL of MCS buffer into the reservoir of the LMC. After rinsing, place a 15 mL conical tube below the column to collect flow through. Apply the whole cell suspension (500 µL) onto the column and let it completely flow through.
   7. Wash the column three times by adding 3 mL of MCS buffer into the reservoir and wait until the column´s reservoir is empty before performing the next washing step.
   8. Collect the unlabeled cells passing the column use for further separation steps (representing the CD45^– fraction). Labeled cells (representing the CD45⁺ fraction) accumulated in the column can be discarded.
6. Accumulate CD31⁺ cells following CD31 microbeads manufacturer’s instructions. For all following steps of CD31⁺ accumulation use MCS buffer.
   NOTE: This step takes about 60 min.
   1. Determine cell number using a Neubauer cell counting chamber (expect about 4 x 10⁶ cells per g muscle tissue).
   2. Centrifuge obtained unlabeled cell suspension from step 3.5 for 10 min at 300 x g at 4°C.
   3. Remove the supernatant completely. Resuspend the pellet in 90 μL MCS buffer and add 10 μL of CD31 microbeads (per 10⁷ total cells or less). Mix the whole suspension and incubate for 15 min in the fridge at 4–8 °C.
   4. Add 1mL MCS buffer and centrifuge at 300 x g for 10 min at 4 °C. Take off the supernatant and resuspend cells in 500 µL of MCS buffer.
   5. For the magnetic cell separation use a medium magnetic column (MMC) with a reservoir volume of 3.5 mL and a capacity of up to 2 x 10⁸ total cells.
   6. Position the MMC in the magnetic field of the separator. Rinse the MMC with 500 µL of MCS buffer. After rinsing, place a 15 mL conical tube below the column to collect flow through.
   7. Apply the whole cell suspension (500 µL) onto the column and let it completely flow through.
   8. Wash the column three times by adding 500 µL of MCS buffer and wait until the column’s reservoir is empty before performing the next washing step. Unlabeled cells passing the column represent the CD45^– CD31^– fraction. Store them for further quality controls.
   9. Remove column from the separator and place it on a suitable collection tube (15 mL tube). Pipette 2 mL MCS buffer onto the column. Immediately flush out the magnetically labeled cells by firmly pushing the plunger into the column. This CD45^– CD31⁺ fraction represents the enriched primary murine EC.
   10. Centrifuge cell suspension for 5 min at 350 x g at 20 °C. Remove supernatant completely and resuspend cells (expect around 0.6 x 10⁶ cells per g muscle tissue) in 1 mL ECM (see step 2.2.). Transfer cells (0.5–0.7 x 10⁶ cells) to a single well of a coated 6-well culture plate (from step 2.3) containing 1 mL of ECM.
7. For cultivation, incubate cells at 37 °C and 5% CO₂ in a sterile incubator. Refresh the ECM every two to three days.

4. Primary Murine MMEC Purification

1. Perform a second cycle of CD31⁺ accumulation, following manufacturer’s instructions (CD31 microbeads mouse protocol; see step 3.6.).
   1. Detach cells with Trypsin/EDTA solution when they are at 80–90% confluence (usually after 7 days). For this, rinse each well with 2 mL sterile PBS, in two consecutive washing steps. Use 800 μL Trypsin/EDTA solution per 6-well and incubate at 37 °C and 5% CO₂ in a sterile incubator for 3–5 min. Stop enzymatic activity by using 1200 μL DMEM containing a minimum of 10% FCS.
   2. Centrifuge the cell suspension for 5 min at 350 x g at 20 °C.
   3. Perform the second CD31-MCS step to increase the purity (according to the CD31 microbeads manufacturer´s instructions; see 3.6.).
   4. Observe cell confluence via bright field or standard phase-contrast microscopy using a
      20x magnification and 0.35 lens numerical aperture. See Figure 1.
      NOTE: Cells can be used for respective experiments or kept in culture by passaging. Use cells for experiments promptly in lower passages. It is not recommended to use cells after passage
      8–10.

5. Quality Control

1. Perform flow cytometry of primary murine MMEC after the second CD31-MCS-step.
   1. Prepare a FACS tube by adding 1 mL flow cytometry (FC) buffer (see enclosed Table of Materials).
   2. Detach cells with Trypsin/EDTA solution when they are at 100% confluence (usually after 14 days). For this, rinse each well with 2 mL sterile PBS, in two consecutive washing steps. Use 800 μL Trypsin/EDTA solution per 6-well and incubate at 37 °C and 5% CO₂ in a sterile incubator for 3–5 min. Stop enzymatic activity by using 1200 μL DMEM containing 10% FCS.
   3. Determine cell number using a Neubauer cell counting chamber and add a minimum of 1 x 10⁵ cells to the prepared FACS tube.
   4. Centrifuge cell suspension for 10 min at 300 x g at 4 °C. Perform two washing steps by removing the supernatant, resuspending cells in 1 mL FC buffer and centrifuging for 10 min at 300 x g at 4 °C.
   5. Prepare a staining mix by adding anti-mouse CD31-FITC antibody (1:100) and anti-mouse CD45-PE (1:100) with FC buffer.
   6. Resuspend cells in 100 μL of staining mix in the FACS tubes. Incubate for 30 min at 4 °C. Add 1 mL of FC buffer. Centrifuge cell suspension for 5 min at 300 x g at 4 °C. Carefully remove supernatant and resuspend cells in 200 μL of FC buffer.
   7. Measure cells in a flow cytometer following the gating strategy is shown in Figure. 2A.
2. Alternatively perform immunofluorescence staining of primary murine MMEC after the second CD31-MCS-step.
   1. Coat the coverslips.
      1. Transfer a sterile 13 mm diameter round glass coverslip into a well of a 24 well plate. Coat coverslip by adding 300 µL of speed coating solution per well. Incubate for 3–5 min at room temperature (RT).
      2. Aspirate and discard coating solution. Rinse each well with 2 mL sterile PBS in two consecutive washing steps. Aspirate and discard PBS.
      3. Seed 1 x 10⁵ cells in 1 mL ECM on the coated coverslip and incubate at 37 °C and 5% CO₂ in a sterile incubator until cells are 99% confluent.
   2. Perform fixation and immunofluorescence staining.
      1. Remove the ECM medium of cultivated cells from 5.2.1.3.
      2. Fix cultured cells by adding 300 µL 4% paraformaldehyde (PFA) with pH of 7.4 per well, for 10 min at 4°C.
         CAUTION: PFA is toxic and must be handled with care.
      3. Perform 3 washing steps by adding 1 mL PBS per well and remove supernatant after 5 min at RT.
      4. Prepare a blocking solution containing 5% BSA, 0.2% Triton-X and 1% goat serum in PBS.
      5. Add 300 µL of blocking solution to each well and incubate for 1 h at RT.
      6. Remove the supernatant and add 200 µL per well of an anti-PECAM-1 (CD31) antibody (1:200) in 5% BSA, 1% goat serum in PBS and incubate at 4 °C overnight.
      7. Perform 3 washing steps by adding 1 mL PBS per well and removing supernatant after 5 min at RT in each step.
      8. Prepare a secondary antibody solution containing Cy3-conjugated anti-rat IgG antibody (1:500) in 1% BSA and PBS. Add 300 µL per well of the secondary antibody solution and incubate for 1 h at RT in the dark.
      9. Perform 3 washing steps by adding 1 mL PBS per well and removing supernatant after 5 min at RT in each step.
   3. Take out the round glass coverslips carefully with forceps and transfer it onto a microscopy slide. Add an appropriate amount of mounting medium containing DAPI on the cell layer and carefully put a cover glass on it (see enclosed Table of Materials).
   4. Observe cells with an appropriate fluorescence microscope equipped with a fluorescence light source (120 W) and two filter sets: An excitation/emission filter set with 550/25 nm and 605/70 nm wavelengths to detect Cy3 540/562 nm.
   5. Use a second filter set with an excitation/emission wavelength of 365/50 nm and 445/50 nm to detect DAPI 350/440 nm. Use a magnification of 40x and 80% intensity of 14.5 mW/cm² for DAPI with an acquisition duration of 30 ms. For detection of Cy3 use 80% of 67.5 mW/cm² with an acquisition duration of 60 ms.
3. Perform quantitative PCR.
   1. Isolation of mRNA.
      1. Follow standard procedures using an acid guanidinium thiocyanate-phenol (AGTP) reagent to isolate mRNA. Use a minimum of 0.5 x 10⁶ cells from CD45^– CD31^– (see step 3.6.8), CD45^– CD31⁺ (see step 3.6.9.) fractions and cultivated primary murine MMEC (4.1.3)
      2. Centrifuge cell suspension for 10 min at 300 x g at 20 °C. Take off the supernatant completely. Resuspend the cell pellet in 500 µL of AGTP reagent and transfer cell suspension into a 2 mL reaction tube. Incubate for 5 min at RT.
      3. Add 100 µL of chloroform and shake vigorously for 15 s. Incubate for 3 min at RT.
      4. Centrifuge suspension for 15 min at 12,000 x g at 4°C.
      5. Take off the upper aqueous phase (containing RNA) carefully and transfer into a 1.5 mL reaction tube. Add 250 µL of isopropanol and incubate for 10 min at RT.
      6. Perform a centrifugation step at 12,000 x g for 10 min at 4 °C.
      7. Take off the supernatant and add 1 mL of 75% ethanol carefully. Centrifuge at 7,500 x g for 5 min at 4 °C.
         NOTE: Pellet is easy to lose.
      8. Remove the supernatant completely. Let the pellet dry on air for 5–10 min.
         NOTE: ethanol needs to be removed but do not dry too much.
      9. Dissolve pellet in 30 µL diethylpyrocarbonate treated water containing in and heat for 10 min at 55 °C.
         NOTE: RNA concentration and purity can be measured by a spectral-photometer.
   2. Perform cDNA synthesis (reverse transcriptase PCR).
      1. Prepare a mastermix containing: 10 µL of PCR buffer (5x), 2,5 µL desoxyribonucleosid-triphosphate (dNTP (10 mM)), 0.5 µL random mixture of single-stranded primer (0.2 µg/ µL), 1 µL of RNase inhibitor (40 U/ µL, 0.4 reverse transcriptase (200 U/µL) and 5.6 µL of diethylpyrocarbonat treated water (see enclosed Table of Materials).
      2. Dilute 500 ng RNA in 30 µL diethylpyrocarbonate treated water in 0.2 mL PCR tubes and add the whole 20 µL mastermix.
      3. Use a PCR cycler performing following steps: 10 min 25 °C, 30 min 50 °C, 5 min 85 °C and hold at 4 °C.
4. Perform Quantitative PCR (qPCR).
   NOTE: Perform qPCR of samples in duplicates or triplicates.
5. Use primer with two different fluorescence dyes. Primer with an absorption of 495 nm and an emission of 517 nm for the gene of interest.
6. For detection of the satellite cell marker gene expression, use primers for paired box protein 7 (Pax7) and M-cadherin (Cdh15) (see enclosed Table of Materials).
7. For detection of the endothelial cell marker gene expression, use primers for claudin-5 (Cld5), occludin (Ocln) and zonula occludens-1 (Tjp1 or ZO1) (see enclosed Table of Materials).
8. As reference gene for 18s rRNA, use a primer with absorption of 538 nm and an emission of 554 nm wavelength.
9. Prepare a qPCR mastermix for each gene of interest with the respective primer. Mastermix contains: 10µl qPCR buffer (2x), 1 µL of Primer for gene of interest (10 µM), 1 µL of Primer for 18s rRNA and 4 µL of nuclease free water.
10. Add 16 µL of the mastermix to a single well of a 96-well reaction plate. Add 4 µL of cDNA (obtained from step 5.3.2.) per well containing the mastermix.
11. Use a real-time PCR cycler performing following steps: holding stage with 50 °C for 2 min and 95 °C for 10 min. Cycling stage with 40 cycles of 95 °C 15 s and 60 °C for 1 min.