Microdialysis of Ethanol During Operant Ethanol Self-administration and Ethanol Determination by Gas Chromatography

1. Stereotaxic Surgery

1. All procedures follow the NIH Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee.
2. Using a stereotaxic apparatus, well-handled Long-Evans rats, anesthetized with isoflurane, are implanted with a 21-gauge cannula (for later microdialysis probe insertion) (Plastics One, Roanoke, VA) above the brain region of interest, and a tether bolt is built into the head stage (used to support microdialysis equipment).
3. Monitor rats carefully during and after surgery. Make sure that all animals receive at least one week of post-surgical care and recovery and are healthy before beginning the following procedures. A JoVE video of the stereotaxic surgery method is available.¹

2. Operant Training

1. After a week of post-surgical recovery, animals are trained to lever press for 10% sucrose solution in a Med Associates operant chamber (MedAssociates, Inc., Vermont, USA) outfitted with a lickometer, retractable lever, and sipper tube (as previously described in Howard et al., 2009).² Software from MedAssociates is used to design all operant programs (MedAssociates, Inc., Vermont, USA).
2. Once trained to respond for sucrose, begin rats on an appropriate training schedule during which ethanol is gradually added into the solution over multiple drinking sessions.
3. For example, our lab currently uses an 8-day training schedule where ethanol is faded into the solution culminating in a 10% ethanol/ 10% sucrose drinking solution. Control animals only receive 10% sucrose or no drinking solution. The specific type of operant schedule to be used can vary widely, but the general procedures described below for microdialysis sampling can still be carried out. ³

3. Pre-microdialysis Procedures: Tethering

1. The night before the second to last training session (in our example this is the 7^th training session), animals are habituated to the tether spring, which attaches to the tether bolt on their head stage. The spring attaches to the microdialysis swivel and counter balance lever arm, which is suspended next to the cage with a ring stand and clamps so that the animal can move freely within its cage. Tethered animals spend the night in the operant room, in their home cage with ad libitum food and water, to habituate them to the tethering set-up.
2. The following day, the rat completes its operant program with the tether in place, to habituate the animal to the feeling of the tether while performing its behavioral tasks. Mount the tethering apparatus to the wall of the operant chamber near the top to allow suspension of the tether and swivel above the center of the roof of the operant chamber. All of this is placed within the sound attenuating chamber. After the session, return the rat to its home cage in the operant room to await microdialysis probe insertion.

4. Pre-microdialysis Procedures: The Microdialysis Probe is Inserted the Day Before the Microdialysis Experiment, after the Rat has Completed Behavioral Training for the Day

1. The day before the microdialysis experiment, after the rat has completed an operant session while tethered, anesthetize the rat with isoflurane and remove the obturator from the guide cannula. Slowly (over ~ 5 min) insert a microdialysis probe, perfused with artificial cerebral spinal fluid (ACSF), through the cranial guide cannula into the brain region of interest. We use laboratory-constructed probes and microdialysis procedures modeled after Doyon et al., 2003, and Pettit and Justice, 1991.^{3, 4}
2. Use the previously discussed tethering apparatus to suspend the inlet and outlet lines of the probe above the animal.
3. Turn the probe flow rate down to 0.2 μl/min. Once again, the rat spends the night in the operant room.

5. Microdialysis Procedures: Collection of Samples During Self-administration Session with Appetitive and Consummatory Phases Separated

1. At least 2 hr before the experiment begins, turn up the probe to its working flow rate. We use either 1.0 or 2.0 μl/min depending on the brain region. Check that the probe flow rate is consistent, and at least 90% of the set flow rate by using a Hamilton syringe to measure volume over time.
2. Dialysis samples (5 or 7 min) are taken before, during, and after the operant session. The sampling interval depends on the brain region, neurotransmitter being analyzed, dialysate concentration of the analyte, and sensitivity of the analytical chemistry equipment to be used for the analysis. The behavioral phases are as follows:
   1. Baseline: At beginning of experiment, collect dialysis samples in the animal’s home cage (4 samples).
   2. Transfer: After collection of home cage baseline samples, transfer the rat to the operant chamber. Transfer of the tethered rat requires extreme care to make sure that the microdialysis fluid transfer line does not become tangled, and that the rat remains calm. Immediately after the transfer, activate the operant program as you change from the baseline/transfer sample to the first wait sample.
   3. Wait: Continue to collect samples as the rat waits for the lever to extend into the chamber (3-4 samples depending on the brain region).
   4. Drink: After the lever is presented and pressed, a bottle of drinking solution (10% sucrose/10% ethanol or 10% sucrose) is made available to the rat for around 20 min (3-4 samples).
   5. Post-drink: After drink period, the bottle retracts, but the rat remains in the operant chamber for around 20 min (3-4 samples).

6. Microdialysis Procedures: Preparation of Microdialysis Sample for Ethanol Analysis

1. Two samples before animals self-administer the ethanol solution, and all samples after, are evaluated for ethanol concentration. Pipette either a 1 or 2 μl aliquot from the sample of interest into a 2 ml glass vial. Then seal the vial with an air tight septum (9 mm Red Poly Screw Cap, PTFE/Sil Septa, Agilent Technologies). The volume of the ethanol analysis aliquot (1 or 2 μl) depends on the total sample volume, and the sample volume required for any additional analyses.
2. If the samples will be used for later neurochemical analysis, store the sample appropriately after pipetting the aliquot for ethanol quantification.
3. For example, our lab analyzes the samples for dopamine. To accomplish this, place the samples on dry ice during the experiment and then, store the samples at -80 °C after the experiment. We use 2 μl aliquots for ethanol analysis of a 5 min sample collected using a flow rate of 2.0 μl/min for probes placed in the nucleus accumbens. This allows at least 7 μl remaining for later analysis of dopamine by high performance liquid chromatography with electrochemical detection.
4. For samples taken from the prefrontal cortex, which has much lower extracellular dopamine concentrations, we use a 1 μl aliquot for ethanol analysis taken from a 7-min sample using a flow rate of 1.0 μl/min.

7. Post-microdialysis Procedures

1. After the conclusion of the microdialysis experiment, anesthetize the rat and remove the probe. Replace the obturator if the animal is not immediately euthanized. Collect the brain within three days of the experiment. Otherwise, visualization of the probe tract may not be possible.
2. The brain should be removed in accordance with approved animal use protocols. We use sodium pentobarbital (150 mg/kg, ip) overdose, followed by cardiac perfusion with saline and then formalin in saline. Submerge brain in a formalin-saline solution for at least 12 hr before sectioning.
3. Coronally section the brain into 100 μm slices. Stain slices with cresyl violet, and examine for correct probe placement.^{5, 6}

8. Analysis of Samples for Ethanol Concentration

1. The collected samples as well as external standards (0.3125 – 20 mM ethanol) are run on our gas chromatograph with flame ionization detection system. This system is comprised of a Varian CP 3,800 gas chromatograph with a flame ionization detector, a Bruker (Varian) 8,400 headspace autosampler heated to 50 °C, and an HP Innowax capillary column (30 m x 0.53 mm x 1.0 μm film thick), with helium as mobile phase.
2. Chromatograms are recorded and analyzed with chromatography software such as the Varian Star Chromatography Workstation software that will be specifically discussed here.
3. To prepare the system for ethanol analysis, heat the autosampler plate using a recirculating water bath, and open the two additional gas tanks (air and hydrogen; helium is always left running to preserve the wax column).
4. Record the gas tank pressures, as well as the number of samples you intend to run so that you can keep count of the number of times the fiber and septum have been used so that they can be changed when appropriate (fiber-500 injections; septum-100 punctures).
5. Initiate the start-up method, which prepares the system to run samples (program parameters in Table 2). Wait for the system to report that it is “Ready”.
6. Initiate a 20-min “burn,” which prepares the fiber for sample analysis by subjecting it to a high temperature to desorb any contaminates it has absorbed while at rest.
7. Standards are made by diluting 238 μl of 95% ethanol with water to a final volume of 100 ml using a volumetric flask in a chemical fume hood. This creates a 40 mM ethanol solution. We use a 1:1 serial dilution to create 20, 10, 5, 2.5, 1.25, 0.625, 0.3125 mM standards. For each concentration of standard, use the same volume aliquot that will be taken from the dialysis samples. Pipette the aliquot into a 2 ml glass vial, and then seal the vial with an air tight septum.
8. All ethanol samples are heated in the autosampler until the entire liquid aliquot has been vaporized. We heat our samples for about 30 min.
9. This section describes the Star Workstation software that we use with our gas chromatograph. Other software may be used, but the description below may not be applicable.
10. To run samples, create a sample list noting which sample is in which autosampler slot, and how many times the sample should be punctured. Be sure to route the data files to your selected folder. Then, select the method of choice for your samples, and begin the run. We use the running parameters noted in Table 2.
11. Our dialysate ethanol program absorbs the sample for 3 min and desorbs into the helium stream, which feeds into the wax column, for 1 min. However, programs can be written to accommodate your specific needs.
12. The sample components separate in the wax column, and then go through the flame ionization detector where the carbon containing compounds, such as ethanol, combust and release ions. This results in increased electron flow from the detector’s anode to the cathode creating a current, which is converted to voltage and recorded resulting in chromatograms like the one shown below (Figure 1). The change in voltage is proportional to the amount of carbon passing through the detector across time.7
13. After the system is finished running the samples you will need to check the integration of each peak. For Star Workstation software, click on the blue peak icon in the tool bar. Click on each peak’s color and drag the arrows to adjust the baseline. Re-integrate each peak before moving onto the next group of peaks. The peak analysis software can be any general chromatography software.
14. For Star Workstation software, click on the batch report icon in your tool bar. Drag each sample from your specified folder to the batch report to print the sample reports. Reporting software can be customized with most common chromatography software systems.
15. The reports can be programmed to show the information of your choice. We currently use peak height, but the reports can also be programmed to use peak area.
16. To shut down the system, initiate the shut-down method (parameters noted in Table 2), turn off the water bath, and close the hydrogen and air tanks when the column oven temperature reaches 30 °C.

9. Ethanol Data

1. Plot the peak height as a function of each known external standard ethanol concentration (Figure 2A). Use the linear function given by these points to calculate the ethanol concentration from the peak height given by each dialysate sample.
2. By plotting the concentration of ethanol in the dialysate over time, we can see the pattern of ethanol levels in the brain region of interest during our behavioral session. Example data, shown below (Figure 2B), are represented as the concentration of ethanol in the dialysate across time during the self-administration session.

10. General Maintenance: The Fiber Should be Changed Every 500 punctures, and the Septum Every 100 Punctures

To change the fiber

1. On the gas chromatograph key pad press Menu, select 8,400, press enter; select change syringe, press enter. The autosampler will position itself to allow the fiber (SPME Fiber Assembly, 75 μm Carboxen-PDMS, Supelco Analytical, Bellefonte, PA) to be removed.
2. Open the door covering the fiber. Unscrew the lock nut, and move the latch to allow the fiber assembly to be removed. Take the fiber assembly out. Unscrew the nut that holds the fiber in the assembly, and then unscrew the fiber.
3. Replace the old fiber with a new one, and reassemble the fiber assembly. Replace the assembly in the autosampler, re-latching and screwing the assembly into place. When you are finished, press “change done” on the key pad, and the autosampler will ready itself for use.

To change the septum

1. First, unscrew the cap covering the septum (Hi-Temp .450 Dia. Generic Conditioned Septa, Varian) and then remove the old septum. Seat the new septum down into the fitting. Re-screw the cap and tighten it with the wrench. Then, use an injection needle to puncture the septum so that the fiber will not break the first time the septum is punctured.

11. Representative Results

Figure 1 shows example chromatograms for three concentrations of ethanol standards and for a rat dialysate sample collected at the end of the ethanol self-administration session. Ethanol peaks should be relatively symmetric, have consistent retention times, and a signal to noise ratio > 10. Failure to meet these criteria means that your system requires maintenance. Quality chromatography and correctly prepared standards produce a linear standard curve (R2 ≥ 0.99; Figure 2A) that is used to calculate the ethanol concentration of dialysate samples collected from ethanol self-administering animals over the course of their self-administration session (Figure 2B).

Figure 1. Example chromatograms. One μl of ethanol standard or dialysate sample was loaded into a gas chromatograph vial and analyzed as described in the text. A) Overlay of peaks generated from 2.5, 5.0 and 10 mM ethanol standards. B) Peak generated from a dialysate sample from a rat that has self-administered ethanol.

Figure 2. Graphical results from example experiment shown in Figure 1. A) Ethanol standard curve. B) Time course of dialysate ethanol concentration across an ethanol self-administration session.

Table 2. Gas chromatograph with flame ionization detection system parameters. This table shows the parameters for the three programs used to prepare (start-up settings), run (running settings) and maintain the system while not in use (resting settings).