Analysis of Gene Expression Changes in the Rat Hippocampus After Deep Brain Stimulation of the Anterior Thalamic Nucleus

NOTE: Ethics Statement: All procedures discussed in this manuscript are in accordance with the NIH guidelines for Animal Research (Guide for the Care and Use of Laboratory Animals) and are approved by the Harvard Medical School IACUC Committee.

1. Pre-surgical Preparation

1. Make sure that all surgical instruments are sterilized by either autoclaving or by cleansing with antiseptic solution and/or ethanol as necessary. Where possible, use sterile disposable equipment such as scalpels, needle and syringes.
2. Cover the workbench with surgical drapes and ensure that there is a biohazard waste disposal available.
3. Weigh the rat and calculate anesthesia dose. Use a Ketamine/Xylazine mix (Ketamine 75 mg/kg and Xylazine 10 mg/kg) to anesthetize the rats.
   NOTE: Between 200-250 g is the optimal weight for proper fixing of the animal in the stereotactic frame as well as for accurate targeting of the ATN. Isoflurane can also be used as the anesthetizing agent.
4. Mount and secure two sterile electrodes (sterilized by autoclaving or with ethylene oxide) on the electrode holder  (Figure 2) of a stereotactic surgical frame and with the help of a microscope (10-40X magnification), inspect the tips of the electrode for proper alignment. Secure the two electrodes 3.0 mm apart.
   NOTE: Take care to avoid damaging the electrode tip by touching on hard surfaces.

2. DBS Surgery

1. Inject the Ketamine/Xylazine mix intraperitoneally to the animal and confirm that the animal has reached a surgical plane of anesthesia (by checking for the toe-pinch reflex, respiratory rate and depth and regularity of breathing). Remove hair from the region of the scalp that will be incised and disinfect the scalp with 3 alternating scrubs each of betadine and either alcohol or sterile saline. Secure and position the animal on the stereotactic frame.
2. Use circulating warm water pads to maintain the animal’s body temperature at an optimal level. Apply eye lubricant to the animal’s eyes to protect from overdrying. 
3. Use the following stereotactic coordinates for targeting the ATN (Anterior thalamic nucleus): anterioposterior -1.6 mm, mediolateral 1.5 mm and dorsoventral 5.2 mm¹.
   NOTE: The stereotactic coordinates are based on the Paxinos and Watson (6^th edition) rat brain atlas³⁶.
4. Make an incision in the scalp sagittally to reveal the skull. Using a pair of retractors secure the incised scalp to expose the skull. Use sterile saline to flush the incision. If incision is too wet, use sterile cotton swabs to dry.  Locate the bregma and mark with a black marker. To guide the position of the burr holes, make two more marks at approximately 1.5 mm mediolaterally on both sides from the sagittal suture and 1.6 mm posterior to the coronal suture.
5. Use a hand-held drill to make the burr holes. Make sure the tip of the burr hole is sterile by sterilizing it with ethanol. Hold the drill at about a 45° angle to the skull surface when drilling. Frequently switch between the two burr holes to avoid excessive heat at the location of any burr hole.
6. Continue drilling until the dura is exposed. Using a needle with its tip bent resembling the shape of an ‘L’, remove any broken pieces of bone that would obstruct the insertion of the electrode. Take care to avoid damaging the underlying dura and/or brain tissue while removing bone fragments using the bent needle.
   NOTE: Using a blunted needle or fine blunt forceps is also an option.
7. Fix the dual electrode assembly to the rotating handle of the stereotactic frame and fix the handle at a 90° angle. Using the adjustments in the stereotactic frame, position the left electrode exactly above the bregma.
8. Using the stereotactic adjustments for mediolateral positioning, precisely move the left electrode 1.5 mm to the left side of bregma such that now there are two electrodes perfectly aligned along the coronal suture but spaced apart at 1.5 mm mediolaterally from the bregma.
9. Using the anterioposterior stereotactic adjustments, move the electrodes 1.6 mm posterior to the coronal suture.
10. Use the dorsoventral adjustments to lower the electrodes to first check if the burr holes have been made at the right location such that the electrodes can be inserted with ease, without touching the rough edges of the burr holes. If so, insert the electrodes to a depth of 5.2 mm from the surface of the skull.
11. Connect the electrodes via leads to a stimulator set at 130 Hz, 2.5 V and 90 µsec pulsewidth¹.
12. Deliver high frequency stimulation for an hour (or for a desired period of time as per experimental setup). During the course of the stimulation, remember to assure proper anesthetic depth regularly by checking for the absence of foot withdrawal in response to a toe pinch. Supplement anesthesia with approximately half the initial dose used to induce anesthesia. Avoid contamination of your sterile gloves when checking anesthetic depth and change them if necessary.  Perform unilateral or bilateral stimulation based on one’s experimental needs. Include controls such as low frequency stimulation (for e.g., 10 Hz) and unstimulated animals (inserting electrodes with no subsequent stimulation).
13. After stimulation is done, remove the electrodes carefully and suture the incision with 3-0 sutures or with sterile surgical staples.
14. Administer buprenorphine (0.05 mg/kg) subcutaneously as analgesia. Monitor the animal until it returns to normal activity and then return it to the housing facility.
15. After a set period of time based on the experimental design (for e.g., 0, 3, 6 or 12 hr post DBS surgery), euthanize the animal with anesthesia overdose. After confirming the absence of vital signs, decapitate the animal.
16. Dissect out the brain by first removing the skin using scissors. Cut through the bone along the sagittal suture using dissection scissors. Make two more incisions (about an inch each) through the bone on both the lateral sides. Using forceps, lift the partially severed piece of bone from the top of the skull to expose the brain.
17. Using fine scissors or forceps, dislodge the brain from the skull and transfer to a Petri dish with cold PBS on ice.
    NOTE: Take care to avoid damage to the brain while making the incisions through the bone.

3. Hippocampus Isolation

NOTE: Perform all the subsequent steps in this section on ice.

1. Place the brain on a pre-cooled acrylic brain matrix on ice. Using a razor blade, cut the brain coronally at approximately 7-8 mm from the anterior-most edge of the brain. Make a second cut coronally and posterior to the first cut such that an approximately 5 mm thick brain slice could be removed.
2. Transfer the brain slice to a Petri dish with ice-cold PBS. Using razor blade, sever the two hemispherical sections and take care to note which hemisphere corresponds to the left and right sides respectively. This is especially important while performing unilateral stimulations.
3. Using fine forceps and scissors remove the hippocampus carefully.
4. Flash freeze the hippocampal tissue on dry ice and store in -20 °C freezer until ready for the subsequent RNA extraction steps.
   NOTE: For long term storage, it is advisable to store tissue in a -80 °C freezer.

4. RNA Extraction and Quantitative PCR

1. Make sure the tissue stays frozen on dry ice until ready for homogenization.
2. NOTE: Perform this step in the hood.
   1. Add 1 ml of Tri reagent to hippocampal tissue in a 1.5 ml centrifuge tube and homogenize it by first pipetting multiple times until the tissue is broken in smaller pieces. Further homogenize the tissue by passing through a syringe with a 25 G needle until there is no unbroken tissue visible. Perform the homogenization steps on ice.
   2. After homogenizing the tissue, allow the cell suspension to stand at room temperature for 5 min to allow cell lysis.
      NOTE: After this point the cell suspension can be quick frozen and stored in -70 °C until further processing.
3. Add 0.2 ml chloroform and mix by vortexing for 20 sec and let the solution rest at room temperature for 15 min.
4. Centrifuge the samples at 12,000 x g for 15 min at 4 °C. After centrifugation, remove the tubes carefully without disturbing the three separate layers that will be visible.
   NOTE: The bottom (red) phase contains proteins, middle cloudy phase contains DNA and the top clear phase contains RNA.
5. Transfer the top clear phase (RNA) into a fresh 1.5 ml centrifuge tube (typically 500 µl of the RNA containing phase is obtained). To this add 0.5 ml isopropanol and mix by vortexing. To this add 2-3 µl glycogen (20 mg/ml) as the carrier for the RNA. Allow the sample to rest on the table at room temperature for 10 min.
6. Centrifuge the sample at 12,000 x g for 1 hour at 4 °C. Make sure that the RNA pellet is visible at the bottom of the tube.
7. Discard the supernatant and wash the RNA pellet by adding 1 ml of 75% ethanol (made with nuclease free water). Invert the sample multiple times until the pellet dislodges from the bottom of the tube and floats in the solution. Centrifuge the solution at 12,000 x g for 10 min at 4 °C.
   NOTE: The pellet in 75% ethanol can be stored at -20 °C until further steps.
8. Allow the RNA pellet to air dry by leaving the tubes open for 5 min at room temperature. Take precaution to avoid over drying the pellet as this might affect its dissolution in water in the subsequent step.

5. Removing DNA from the RNA Preparation

1. Add 9 µl of nuclease free water to the RNA pellet and make sure the pellet is dissolved before proceeding by gently vortexing the tube. To this add 1 µl of 10x Dnase I Buffer and 1 µl recombinant DNase (from DNase I kit), mix by gently flicking the tubes, briefly spin the tubes and incubate at 37 °C in a water bath for 30 min.
2. Add 2 µl DNase inactivation reagent (from DNase I kit) to the sample and incubate at room temperature for 2 min and mix often by gently flicking the tube. Centrifuge at 12,000 x g for 10 min and transfer approximately 10 µl of the clear supernatant to a fresh 1.5 ml centrifuge tube.
3. Measure RNA concentration using a nanodrop or spectrophotometer.

6. Making cDNA from RNA

1. Add requisite volume that contains 1 µg of RNA and bring the total volume to 8 µl by adding nuclease free water. To this add 1 µl 10 mM dNTP mix and 1 µl of random hexamers from the Superscript First Strand Synthesis Kit. Gently mix and incubate at 65 °C for 5 min in either a water bath or on a pre-programmed thermocycler.
2. Make a premix containing the following reagents: 2 µl of 10x RT Buffer, 4 µl of 25 M MgCl₂, 2 µl of 0.1 M DTT and 1 µl of RNAse OUT (40 U/µl).
   NOTE: Volumes given are per reaction, user would need to scale up according to one’s experimental needs.
3. After removing the RNA containing sample from 65 °C, set the tube at room temperature for a minute. Add 9 µl premix solution and incubate at room temperature for 2 min. Add 1 µl of Superscript II reverse transcriptase enzyme (50 U/µl) and then incubate at room temperature for 10 min.
4. Incubate the samples at 42 °C for 50 min for the reverse transcription to occur.
5. Incubate the samples at 70 °C for 15 min to terminate the reaction.
6. Place the samples on ice briefly and add 1 µl RNAseH (2 U/µl) and incubate at 37 °C for 20 min.
7. Briefly centrifuge the tubes at 3,000 x g to spin down the condensate liquid.
   NOTE: This is the cDNA that will be used for quantitative PCR. cDNA can be stored in -20 °C freezer until PCR setup. cDNA can also be diluted with nuclease free water before proceeding to PCR.

7. Quantitative PCR

1. Make a master mix containing the following reagents (volumes given are per reaction):6.3 µl of 2x Sybr Green Mix, 0.6 µl of Forward Primer (10 µM), 0.6 µl of Reverse Primer (10 µM) and 0.5 µl of Nuclease free water.
   NOTE: Primer design was done using the ‘Pick Primers’ option in NCBI’s nucleotide sequence page for the gene of interest.
2. Add 10 µl mastermix to each well of a 96 well- PCR Plate. Add 2.5 µl of cDNA made earlier to the well containing the mastermix. Make sure to set up triplicate PCR reactions for each sample.
3. After completing the addition of mastermix and cDNA, cover the PCR plate using an optical adhesive sheet to seal the wells. Spin the plate briefly for 5 min at 500 x g in a tabletop centrifuge to settle all the liquid to the bottom of the well.
4. Load the PCR plate onto a pre-programmed RT-PCR machine. Use the PCR parameters provided in Table 1.
5. Data analysis: Use the C_t values from the qPCR output for further calculations. Along with the test gene, set up PCR reactions for internal control genes, 18S rRNA (to ensure equal input) and β-actin (negative control). Calculate fold changes based on ΔΔCt method ³⁷.