Combining Laser Capture Microdissection and Microfluidic qPCR to Analyze Transcriptional Profiles of Single Cells: A Systems Biology Approach to Opioid Dependence

This study was carried out in accordance with the recommendations of Animal Care and Use Committee (IACUC) of Thomas Jefferson University and Drexel University College of Medicine. The protocol was approved by the Thomas Jefferson University and Drexel University College of Medicine IACUC.

1. Animal model

1. Insert two 75 mg slow-release morphine sulfate pellets or two placebo pellets subcutaneously in adult male Sprague-Dawley rats.
   1. Use a gown and gloves appropriately for minor sterile surgery. Shave the rat dorsum with clippers if necessary.
   2. Apply vet ointment to the animal's eyes. Anesthetize the rat with approximately 20 s of isoflurane inhalation. Anesthesia is confirmed by loss of consciousness.
   3. Make a midline incision in the rat dorsum with bead-sterilized blunt scissors and separate the dermis from the body wall with a bead-sterilized probe. Insert the pellets under the dermis with bead-sterilized forceps. Suture the incision closed with a sterile needle.
      NOTE: The entire procedure takes about 5 min per rat. Fresh sterile gloves are used for each rat.
   4. Place the rat into an isolation cage for postsurgical recovery. Check for a heartbeat and regular respiratory rhythm. Observe the rat until consciousness is regained. Assess for postsurgical pain.
   5. Assess the rats 8 h postsurgery and every 12 h after for recovery and infection. Place rats in a cage with the rest of the cohort when they are fully recovered from surgery, about 24 h postsurgery.
2. Give an intraperitoneal naltrexone injection (75 mg/kg) to the G and the Withdrawal cohorts following 6 days of morphine exposure.
   NOTE: There were four rat cohorts in this representative experiment (see Figure 1).

2. Sample harvesting

1. Harvest the brain 6 days following the pellet insertion or 24 h following the naltrexone injection.
   1. Place the animal in an isoflurane chamber for approximately 30 s or until loss of consciousness occurs, indicated by a lack of motion and decreased respiratory rate.
   2. Use a sharp guillotine to rapidly decapitate the animal.
   3. Dissect the brain out from the animal's skull.
   4. Use a sharp handheld razor to make the following gross incisions to the removed brain: First, slice off the cerebellum and discard. Second, separate the brainstem from the forebrain with a transverse incision. Third, hemisect the forebrain and/or brainstem with a midline sagittal incision.
   5. Place the forebrain and brainstem into a plastic tissue-embedding mold with 3-4 cm of Optimal Cutting Temperature compound (OCT) in the bottom of the mold. Cover the rest of the sample with OCT.
   6. Immediately place the plastic tissue-embedding mold with the sample covered with OCT into a bath containing dry ice and methanol. Do not let the methanol spill into the tissue-embedding mold. Keep the embedding mold with the brain sample in the methanol-ice bath until tissue collection is finished (~10-15 min maximum).
   7. Place the brain sample into a -80 °C freezer as soon as possible.
2. Harvest the gut samples concurrently.
   1. Following rapid decapitation, make a midline incision in the animal's abdomen with a scalpel.
   2. Find the cecum and sever its connection to the ascending colon.
   3. Squeeze the cecal contents into a 15 mL conical tube.
   4. Immediately place the conical tube on dry ice and put into a -80 °C freezer as soon as possible.
      NOTE: The small intestine contents can also be collected using the same methods as a negative control.

3. Slicing

1. Slice the hemisected rat forebrain using a cryostat.
   1. Remove the plastic embedding mold with the forebrain from the -80 °C freezer and place into a -20 °C cryostat.
   2. Remove the OCT-embedded hemisected forebrain sample from the embedding mold. Use a razor to slice the corners of the plastic embedding mold vertically if necessary. Mount the forebrain for rostral to caudal coronal slicing on a cryostat chuck using OCT.
      NOTE: Anatomic landmarks to identify the CeA include the optic tract and stria terminalis (Figure 2B). The optic tract branches from the optic chiasm and tracks dorsal-lateral as the brain is sliced rostral to caudal. When the optic tract has a morphology similar to what is seen in a rat brain atlas bregma -2.12 mm¹⁰, test slices may be viewed under a microscope. The optic tract and stria terminalis morphology can be checked in a rat brain atlas¹⁰ to identify the bregma and whether the CeA surrounds the stria terminals.
   3. Slice 10 µm thick coronal sections from the hemisected forebrain rostral to caudal until sections containing the CeA are reached.
      NOTE: The width and height of the sections are approximately 200 mm.
   4. Collect 10 µm sections containing the CeA, or the preferred brain region, by thaw-mounting 10 µm sections onto plain glass slides. Immediately place the glass slides onto a metal pan resting on dry ice. Put the slides with brain sections into a -80 °C freezer as soon as possible.
      NOTE: Multiple slices may be placed on the same slide. If using a different cell type stain for slices on the same slide, leave about 100 mm between slices so a hydrophobic pen can be used to separate cell type-specific antibody solutions on the slide. Leave about 20 mm from the edge of the slide on each side of the slice.

4. Immunofluorescence staining

1. Stain the forebrain sections for the brain cell of choice (e.g., neuron, microglia, astrocyte, etc.) using immunofluorescence.
   1. Remove one or more slides with 10 µm sections of the CeA from the -80 °C freezer.
   2. Fix the slides with 75% ethanol for 30 s. Remove the excess liquid.
   3. Block the slices for 30 s with 2% bovine serum antigen (BSA) in 1x phosphate buffer saline (PBS). Wash 1x with PBS.
   4. Add a primary antibody solution to the slide for 2 min. Wash 1x with 2% BSA solution.
      NOTE: The primary antibody solution is composed of 2% primary antibody, 1% RNase Out, and 96% of same BSA PBS solution for the blocking step above (step 4.1.3). The representative experiment used an anti-NeuN antibody, an anti-Cd11β antibody, and an anti-GFAP antibody in the following quantities: 3 µL of primary, 1.88 µL of RNA inhibitor, and 145.12 µL of BSA solution.
   5. Add the secondary antibody solution to the slide for 3 min. Wash 1x with PBS.
      NOTE: The secondary antibody solution is composed of 1 µL of goat anti-mouse 488 nm fluorescent tag (1:500), 2.5 µL of RNA inhibitor, 1.3 µL of DAPI (1:10,000), and 196.5 µL of 2% BSA.

5. Ethanol and xylene dehydration series

1. Dip the slides into 75% ethanol for 30 s. Immediately after, dip the slides in 95% ethanol for 30 s. Immediately after, dip the slides into 100% ethanol for 30 s. Immediately after, dip the slides into a second container containing 100% ethanol for 30 s.
2. Following the ethanol dehydration series, dip the slides into freshly poured xylene for 1 min. Immediately after, dip the slides into a second container of xylene for 4 min.
3. Remove the slides from the xylene bath and let air dry in the dark for 5 min.
4. Place the slides in a desiccator for 5 min to dry further.

6. Laser capture microdissection

1. If stained, place the slide into the microscope and find the region of interest (CeA) using anatomic landmarks (i.e., the optic chiasm and stria terminalis).
2. Use fluorescence to identify the stained cell type and its nucleus in the region of interest. Choose one cell or multiple cells if doing single cell pooled samples. Mark the cells of interest using LCM software.
3. Place the LCM cap on top of the slice on the region of interest.
4. Use test shots of an infrared (IR) laser to adjust the IR laser strength, size, and duration so that the LCM cap adhesive melts only over the area of the selected single cell. This ensures that no other cells will be collected other than the cells selected.
   NOTE: In this representative experiment, 10 cell pools of the same cell type were used as a single sample to limit the cell-to-cell variability in gene expression between samples with the same treatment. However, this method can be used for true single cell experiments^1,3.
5. Select the individual cells to be collected for analysis using the LCM software tools (Figure 2C). Cells selected must be in the anatomic area of the CeA (or the brain region of choice) based on the rat brain atlas and the bregma¹⁰. Cells should be at least 3 µm from the adjacent stained nuclei.
6. Fire the IR laser to collect the identified single cells.
7. Place the cap in the quality control (QC) station and view it to ensure that only the desired cells were selected. If other cells were mistakenly selected, an ultraviolet laser can be used to destroy the unwanted cells while the cap remains in the QC station.
8. Take a photo of the tissue section from where the cell was collected to document its anatomic specificity. Record the distance of the slice from the bregma if appropriate using a rat brain atlas as a reference¹⁰.
9. Remove the LCM cap from the QC station, attach the sample extraction device, and pipette 5.5 µL of lysis buffer onto the sample.
   NOTE: The lysis buffer solution consists of 0.5 µL of lysis enhancer and 5 µL of resuspension buffer.
10. Fit the ExtracSure device onto a 0.5 mL microcentrifuge tube and place on a hotplate at 75 °C for 15 min.
11. Spin down the sample and lysis buffer for 30 s at low speed (0.01-0.02 x g) and place the collected sample into a -80 °C freezer.

7. Single-cell microfluidic RT-qPCR

1. Preamplification of single cell mRNA for 96.96 Dynamic Array Chip
   1. Combine the forward and reverse mRNA qPCR gene primers for all the genes being assayed in a primer pool for preamplification (500 nM concentration each primer). For example, 1 µL of 100 µM primers in 80 µL plus 120 µL of DNA Suspension Buffer.
      NOTE: The primer sequences used for the representative experiment can be found in O'Sullivan et al.⁴.
   2. In a new 96 x 96 PCR plate, add 1 µL of 5x VILO to each well.
   3. Remove LCM single cell samples from the samples stored at -80 °C, let thaw briefly, centrifuge briefly at a low speed (0.01-0.02 x g), and add 5.5 µL of the lysed single-cell sample to the PCR plate. Each sample is added to its own well.
   4. Place the PCR plate with the samples and VILO into the thermocycler and heat at 65 °C for 1.5 min. Spin the plate for 1 min at 1,300 x g at 4 °C and place the plate on ice.
   5. Add 0.15 µL of 10x cDNA synthesis master mix, 0.12 µL T4 Gene 32 protein, and 0.73 µL of DNA suspension buffer to each well.
   6. Place the PCR plate into the thermocycler and run the following protocol: 25 °C for 5 min, 50 °C for 30 min, 55 °C for 25 min, 60 °C for 5 min, 70 °C for 10 min, 4 °C to end.
   7. Add 7.5 µL of Taq polymerase master mix to each well.
   8. Add 1.5 µL of the primer pool (see above) to each well.
   9. Place the PCR plate in the thermocycler and run the following preamplification protocol: 95 °C for 10 min, followed by 22 cycles of 96 °C for 5 sec, 60 °C for 4 min.
   10. Add 0.6 µL of exonuclease I reaction buffer 10x, 1.2 µL exonuclease I, and 4.2 µL of DNA suspension buffer to each well.
   11. Place the PCR plate in the thermocycler and run the following protocol: 37 °C for 30 min, 80 °C for 15 min.
   12. Add 54 µL of TE buffer to each well. Spin the PCR plate at 1,300 x g at 5 min. Store at 4 °C if immediately continuing to next step. Store at -20 °C if waiting more than 12 h for next step.
2. Prepare the sample plate for the 96.96 Dynamic Array Chip.
   1. In a new 96 well PCR plate, add 0.45 µL of 20x DNA binding dye and 4.55 µL of low ROX mastermix to each well.
   2. Add 3 µL of preamplified sample to each well, spin the PCR plate at 1,300 x g, then put the plate on ice.
3. Prepare the assay plate for the 96.96 Dynamic Array Chip.
   1. In a new 96 well PCR plate, add 3.75 µL of 2x GE assay loading reagent and 1.25 µL of DNA suspension buffer to each well.
   2. Add 2.5 µL of 10 µM qPCR primer to each corresponding well. Spin the PCR plate at 1,300 x g for 5 min.
4. Load and run the 96.96 Dynamic Array Chip.
   1. Prime the chip with control line fluid.
   2. Place the chip in an IFC Controller HX and run the Prime (136X) script.
   3. Add 6 µL of the sample from the PCR sample plate into the corresponding sample wells in the 96.96 Dynamic Array Chip.
   4. Add 6 µL of the sample from the PCR assay plate into the corresponding assay wells in the 96.96 Dynamic Array Chip.
   5. Use needles to pop any air bubbles in the wells of the 96.96 Dynamic Array Chip.
   6. Place the 96.96 Dynamic Array Chip into the IFC Controller HX and run the Load Mix (136x) script.
   7. Remove the chip from the IFC Controller HX, peel off the protective sticker, and place the 96.96 Dynamic Array Chip into a microfluidic RT-qPCR platform. Run the GE Fast 96 x 96 PCR protocol (30 cycles).
      NOTE: The RNA quality and validity of the results are assessed by multiple methods, including assay validation via gel electrophoresis, melting temperature curves, sample and assay replicates, and standard dilution series plots. Additionally, transcriptional findings can be validated by independent methods on brain hemisection including Western blot and immunofluorescence assays.

8. Measuring the bacterial abundance with microfluidic RT-qPCR

1. Extract the bacterial DNA following the directions of the stool DNA extraction kit.
2. Estimate the bacterial DNA concentration using qPCR.
3. Add the extracted bacterial DNA to a new PCR plate. Add 1 µL of extracted bacterial DNA and 9 µL of DNA Suspension buffer.
4. Prepare the assay plate for the 48.48 Dynamic Array Chip (see steps 7.2.1-7.2.2)
5. Prepare the sample plate for the 48.48 Dynamic Array Chip (see steps 7.3.1-7.3.2)
   1. In a new 96 well PCR plate, add 0.45 µL of 20x DNA binding dye and 4.55 µL of low ROX mastermix to 48 wells.
   2. Add 3 µL of the sample from the PCR plate containing the bacterial DNA to the 48 wells and spin down the PCR plate at 1,300 x g for 5 min. Store at 4 °C.
6. Load and run the 48.48 Dynamic Array Chip (see steps 7.4.1-7.4.7).