使用的跟踪经典条件反射，评估胎儿酒精谱系障碍大鼠模型海马功能障碍

摘要

痕量经典条件反射 (ECC) 用于评估了的成年大鼠海马依赖联想学习管理高 (11.9 %v / v) 乙醇浓度早期新生儿大脑发育。ECC 程序一般声音诊断工具检测脑功能障碍跨越心理和生物医学的许多设置。

摘要

新生大鼠进行问卷调查期间产后天 4-9，相对较高浓度的乙醇 （酒精） (11.9 %v / v) 一段时间当胎儿的大脑经历了快速的组织变革是类似于加速大脑在人类妊娠期间发生的变化。这个型号的胎儿酒精谱系障碍 (FASDs) 产生严重的脑损伤，模仿的数量和模式的酗酒，发生在一些酒精孕妇。我们描述了使用的跟踪经典条件反射 (ECC)，高阶 variant 类型的联想学习，以评估长期海马的功能障碍，通常看到酒精暴露的成年子女。在 90 日龄，啮齿类动物是手术准备好录音和刺激电极，测量肌电图 (EMG) 眨眼从左的眼睑肌肉的活动，并分别交付轻微冲击后的左眼。经过 5 天恢复时间，他们经历了跟踪确定酒精暴露与控制大鼠联想学习差异的 ECC 6 各届会议。痕量 ECC 是可以轻松地修改使用相同的设备和软件，以便可以评估不同的神经系统的许多可能的 ECC 程序之一。ECC 程序一般情况下，可以被用于作为诊断工具检测神经病理学侮辱大脑的不同条件，不同的大脑系统。

引言

It is quite hard to imagine that in today's day and age with better health care and access to health services, alcohol abuse remains a major global health concern. Unfortunately, it has been shown that an expectant mother who drinks a high amount of alcohol can have a child with severe brain damage and neurodevelopmental disorders that last a lifetime, as evident in those afflicted with fetal alcohol syndrome (FAS)^1,2,3. In women with some confirmed history of maternal alcohol use, the developing fetus is also susceptible to small amounts of alcohol or different patterns of alcohol consumption that produce varying differences in blood alcohol concentrations. In this latter case, while the children may not exhibit the severe morphological or neurobehavioral disruptions as those with FAS, they may still exhibit lifelong cognitive disabilities and emotional disturbances that range from mild to severe^3,4. Altogether, FAS and less severe forms of prenatal alcohol-mediated disruptions constitute a collection of fetal alcohol spectrum disorders (FASDs). It is no surprise that FASDs are completely preventable, but astonishingly estimates show that in populations where alcohol abuse is quite common, they remain the primary non-genetic cause of neural and cognitive disability, affecting about 2% to 5% of young US children and those in European countries such as France and Sweden. With respect to the incidence of FAS alone within the US, the prevalence is 2 to 7 per 1,000 live births⁵, implying that the overall incidence of FASDs to be much higher than that for FAS.

Neuroimaging studies conducted in children with FASDs have shown that they display brain abnormalities, such as a thinner corpus callosum⁶, smaller anterior cerebellar vermis⁷, and smaller hippocampus⁸. These brain abnormalities underlie some of the long-term neurocognitive disruptions observed in children with FASDs. The exact links that tie variations in maternal alcohol-mediated brain changes and variations in the profile (i.e., type, extent) of particular neurocognitive impairments have yet to be clearly determined. But as a starting point, the hippocampus is an excellent candidate for determining its susceptibility to prenatal alcohol effects. Indeed, children with FASDs exhibit deficits in hippocampal-mediated behaviors such as place learning^9,10 and delayed object recall¹¹.

Rodent models of FASDs have proven to be invaluable in elucidating the mechanisms leading to neurocognitive disruptions seen in children with FASDs. A well-established binge-exposure model that we have adopted involves delivering alcohol to rats during postnatal days 4-9^12,13, a period when the brain undergoes rapid synapse and dendritic contact formation, comparable to human fetal week 24 and extending into the 3^rd trimester^14,15,16,17. This particular model induces significant loss of hippocampal neurons^18,19 and neurons in many other brain regions such as the cerebellum^{12,13,14,15,16,17,18,19,20,21,22,23}, accompanied by severe impairments in cognitive functions spanning different domains^21,24,25. Cognitive disruption from early alcohol exposure in rats may be assessed in different ways, particularly with eyeblink classical conditioning (ECC). ECC is a paradigm that has been utilized for more than a century to scientifically investigate the fundamental basis of learning^26,27 and as such, provides a useful method to better understanding the adverse neurocognitive consequences resulting from fetal alcohol exposure. It is a very flexible paradigm that allows investigators to use a variety of different ECC procedures, any of which can be examined across many mammalian species ascending the phylogenetic scale (from mice to humans) and over different courses of brain development^28,29,30,31. Furthermore, the fundamental neural circuits that mediate associative learning in this paradigm are supported by experimental and neuropsychological reports in these same species^{26,32,33,34,35,36,37}.

One form of ECC, trace ECC is demonstrated in this paper (Figure 1). To provide context, it is compared against the more traditional form - delay ECC. The ECC paradigm was modeled after classical conditioning using dogs, first carried out by the Nobel-Prize winning physiologist, Ivan Pavlov. Pavlov discovered that certain stimuli such as tones do not naturally elicit salivation, but when it precedes and overlaps with the delivery of food, the salivary response can be strengthened from repeated presentations of the two, provided that this tone-food contingency is maintained. This is an example of delay ECC, with the notion that associative strength is mediated by immediate temporal contiguity between the two stimuli, thus making learning conditions optimal for an animal. He also tested other variations of the tone-food contingency, such as turning the tone off and leaving a "trace" period before delivering the food. When these two stimuli were discontiguous enough, it became much harder for the dogs to emit salivation responses prior to the delivery of the food. The discontiguity between the tone being turned off and the delivery of the food is thus an example of trace ECC. As rodents do not naturally salivate to the presence of food, more species-relevant stimuli such as mild shock are used instead; they also do not naturally emit defensive eyeblink responses to tones. With this backdrop, rodent ECC procedures involve presenting a tone at a given decibel level and pairing it in some fashion with mild shock to either the eyelid muscle (orbicularis oculi) or the temporalis muscle to elicit an eyeblink response. The tone is considered a conditioned stimulus (CS) while the shock is considered an unconditioned stimulus (US). In delay ECC, the CS is presented first; this stimulus remains on for a given duration. Afterwards, the US is delivered. These two stimuli overlap for a given duration, and then both terminate simultaneously; the resultant eyeblink response emitted due to the US is considered an unconditioned response (UR). In this procedure, rodents learn to emit eyeblink responses sometime after the CS is presented, but just before the US, in order to anticipate this aversive stimulus. The learned eyeblink response is referred to as a conditioned response (CR). For trace ECC, the CS and US are separated by a period of time that is void of stimuli known as a trace interval; they do not overlap in time as in delay ECC. During this interval, the animal is tasked to resolve the associational requirements between stimuli. Similar to delay ECC, learning occurs when the animal consistently emits a blink response after the CS turns off, but immediately before delivery of the US. Over some amount of acquisition training (CS paired with US), learning curves (i.e., based on different CR measurements) develop. Lesion and neuroimaging studies show that successful learning in delay ECC is dependent on having intact cerebellar-brain stem neuro-circuitry^38,39,40, whereas trace ECC is a higher-order procedure that requires additional neural engagement from the hippocampus^41,42,43,44 and other cortical structures^45,46. Because of the timing-related requirements needed in order to acquire trace CRs successfully, this task is also more difficult to learn (even for normal subjects).

Figure 1: Trace eyeblink classical conditioning. An actual waveform is shown that is representative of an adult rat in the unintubated-control (UC) group. The tone CS (85 dB, 2.8 kHz) is first presented for 380 ms. A trace interval of 500 ms ensues, where no stimuli are present. Afterwards a shock US (1.6 mA) is delivered for 100 ms. Successful learning in this task occurs when the frequency (%) or amplitude (in volts) of eyeblinks during the conditioned response (CR) time window (Total CR period) increases over many sessions of training. In particular, rodents with an intact hippocampus will usually emit more well-timed CRs (Adaptive CRs) just prior to the onset of the shock US (within a 200-ms window). Startle responses (SRs) during the first 80 ms after tone CS onset and unconditioned responses (URs) are also measured. Non-associative SRs are typically low or nonexistent in well-trained rodents, while URs are expected to be high in frequency and amplitude. This task requires that the rodent learn to bridge the association between the CS offset and US onset (during the trace interval), therefore making it inherently more difficult to acquire compared to delay ECC. Please click here to view a larger version of this figure.

Here we demonstrate the adverse functional consequences of neonatal alcohol exposure that is delivered in a binge-like manner, as assessed by a trace ECC procedure that delivers an 85 dB tone CS (2.8 kHz) which remains on for 380 ms, followed by a 1.6 mA shock US which remains on for 100 ms, and these stimuli are separated by a trace period of 500 ms. We have reported on the utility of this behavioral assay in previous studies examining choline intervention and iron supplementation in mitigating the effects of neonatal alcohol exposure^18,47. Indeed, trace ECC can be used as a diagnostic tool to assess neonatal alcohol-induced hippocampal pathology. The advantage it has over delay ECC is that it is more sensitive to detecting disturbances in hippocampal function, which is compromised in humans with FASDs.

Demonstration of ECC extends far outside the fetal alcohol field. Many variants of ECC (e.g., delay, trace, compound, reversal) can be used to elucidate ontogenetic differences in learning across development, the neurobiological basis of associative learning in normal mammals, as well as the vulnerabilities of different brain systems to many challenges, including (but not limited to) teratogens, environmental toxins, traumatic brain injury, neurodegenerative diseases, and psychiatric conditions.

研究方案

NOTE: All procedures were approved and carried out in accordance with the policies set forth by the East Carolina University IACUC. Long-Evans rats were generated from females mated with male breeders. Pups from all three treatment groups (see 1.1) were generated within a litter from the same dam. Five litters were produced and each litter was culled to 8 pups on postnatal day (PD) 3. The remaining two pups from each litter were assigned to separate experiments. Both male and female offspring (one per exposure group) were included in the study. A total of N = 27 adult offspring were examined in this study; 3 rats were excluded due to broken 3T wire leads (see 3.1.1) which were irreparable on Day 1 of ECC training.

1. Preparation of Groups, Materials, and Solutions

1. On PD 3, remove pups from the dam and place them on a thermo-regulated water heating pad to keep warm. Use a random selection procedure to place rat pups into one of three groups: (1) alcohol-intubated (AI); (2) sham-intubated control (SI); or (3) unintubated-control (UC). Carry out approved procedures for identification, such as [non-toxic] ink tattooing of their paws with a 30G x 1/2 in needle following a numbering scheme. Return the pups to the dam when finished with tattooing.
2. Prepare an intragastric intubation tube(s) using polyethylene (PE) size 50 and 10. The overall length of this tube is user-dependent and can be adjusted accordingly.
   1. Use micro dissecting scissors to cut a 3 in piece of PE-50 tubing and a 6 in piece of PE-10 tubing.
   2. Carefully attach the PE-50 tubing to a sterile 22 G x 1 in hypodermic needle.
   3. Insert one end of the PE-10 tubing into the open end of the PE-50 tubing. A diagonal cut may be needed at the tip of the PE-10 tubing in order to direct it into the PE-50 tubing.
   4. Cut a small piece of PE-50 tubing (about 2-3 mm) and insert it on the open end of the PE-10 tube. This will serve as a visual stopper.
   5. Store the complete intragastric intubation tube in fresh 70% isopropyl alcohol to keep it sterile.
3. Carry out aseptic procedures to prepare a stock milk solution in 50 mL bottles. The stock milk solution is based on a diet formula first established by West et al. (1984)⁴⁸. Store the bottles at -18 °C until time of use. Thaw two bottles of milk using warm water prior to use.
   1. Draw out 7.16 mL of 95% ethyl alcohol (USP) using a sterile syringe and add it to one 50 mL milk bottle. This makes an 11.9% v/v alcohol solution that is delivered to rat pups over the first two feedings of the day (see 2.1.1); the total daily dose of alcohol is 5.25 g/kg/day. It is designed to deliver the alcohol in a volume (0.0278 mL/g per feeding) in enriched milk solution.
      NOTE: This amount can be adjusted based on litter size and expected usage over 3-4 days. The amount of alcohol added must also be adjusted in order to make an 11.9% v/v solution.
   2. Shake this bottle well and label it accordingly (i.e., 11.9% v/v Ethyl Alcohol in Milk).
   3. Label the second bottle of milk (without alcohol added) accordingly (e.g., Milk-Only) and use it for supplemental feedings as indicated in 2.1.1.
   4. Store both milk bottles in the refrigerator.

2. Neonatal Alcohol Exposure (Postnatal Days 4-9)

1. Give AI and SI pups 4 total intubations per day starting on PD 4. Separate each intubation by 2 h. Because AI pups are intubated with an actual solution, each of their intubations are considered feedings.
   1. Intubate the AI pups with the 11.9% v/v Ethyl Alcohol in Milk solution during the first two feedings of the day, and then intubate them with the Milk-Only solution during the last two feedings of the day to supplement their growth.
   2. Intubate the SI pups 4 times without any solution every two hours, ensuring the same duration of PE-10 tube exposure that AI pups endure. The UC group does not receive any intubations.
2. Remove pups from the dam and place them on a thermo-regulated water heating pad to keep warm. Weigh each pup and record its body weight (in g). Refer to a pre-made feeding chart to determine the intubation volume for each AI pup and note it in a record sheet.
   1. Place the 11.9% v/v Ethyl Alcohol in Milk solution in a warm water bath and shake it well.
   2. Intragastric intubation procedure:
      1. Flush out the intragastric intubation tube that was stored in 70% isopropyl alcohol well with warm water.
      2. Extract the correct amount of solution using a sterile 1 mL syringe.
      3. Dip the tip of the intubation tube (PE-10 end) in fresh corn oil (this facilitates insertion).
      4. Measure the length of the PE-10 tube from the pup's mouth to its stomach. Adjust the PE-50 stopper to guide with the stopping point.
      5. Carefully insert the PE-10 tube into the pup's mouth, proceeding down its esophagus, slightly passing through the gastroesophageal sphincter, and into its stomach. Examine the pup, stabilize it, and depress the syringe plunger to deliver the solution. Do this at a slow rate.
         NOTE: To avoid accidentally inserting the tube into the trachea, first direct the tube so that it curves and makes contact with the soft palate prior to proceeding forward. Use the palate as a guide, as the tube will naturally deflect off this region and be guided down the esophagus. Otherwise, any substantial deviation of the tip ventrally may cause it to enter into the trachea. It is best to remove the tube and not to proceed with the intubation if one is not certain about its entry position - reinsert the tube correctly on the next attempt.
      6. Carefully remove the PE-10 tubing and examine the pup for backwash of solution, blood, or physical injury. Replace pup with its littermates if it is fine. The entire intubation procedure takes approximately less than 1 min for each pup and approximately 4 min total for the 4 intubation-treated pups in each litter (2 AI, 2 SI).
      7. Intragastric intubation procedure for SI pups: Follow the same PE-10 insertion procedure as that for AI pups, but without any solution for the same duration (1 min).
      8. Return all pups back to the dam immediately after completing procedures for the last pup in the litter. Return the dam to the vivarium until the next feeding session (in 2 h).
      9. Flush out the intragastric intubation tube with warm sterile water and replace it in 70% isopropyl alcohol for storage. Flush the tube well with water prior to each feeding session.
      10. Repeat 2.2 (except weighing pups) to 2.2.2.9 for the second feeding session. At the last two feedings, intubate the AI pups with the Milk-Only solution using the same intubation volumes determined in 2.2.
3. Weigh the rats at regular intervals such as on PD 15 (when eyes open), PD 21 (weaning), PD 30, PD 60, and PD 90 (surgery) to obtain representative growth curves.

3. Fabrication and Modification of Electrodes

1. Fabricate one electromyographic (EMG) "headstage" for each rat (Figure 2). The headstage allows for recording of eyeblink responses via the eyeblink system (Figure 4).
   1. Construct a headstage that consists of two size 3T polytetrafluoroethylene (PTFE)-coated stainless steel wires (5 cm each), one size 10T PTFE-coated stainless steel wire (5 cm), three male contact pins, and one micro strip socket insulator that is cut-down to 3 holes (see 3.1.4).
   2. Strip off the PTFE coating from the 10T wire by grasping the center with a serrated high precision tweezer and removing the coating in both directions with the smooth high precision tweezer. Ensure that all traces of coating have been removed from this wire.
      1. Crimp one end of the 10T wire to a contact pin with a serrated platform dental plier; this will serve as a ground wire. Hold a 3T wire carefully with the smooth high precision tweezer 1 to 1.5 mm from one end. Strip off the 1 to 1.5 mm PTFE coating while leaving the rest of the coating intact. Crimp a contact pin to the exposed end of the 3T wire.
      2. Carry out the same procedures for the second 3T wire. Both wires will serve as the positive and negative leads of the EMG recording electrodes.
   3. Perform tug tests on all wires to ensure that they are secured to the pins. Take caution not to bend or damage the wires.
   4. Cut a segment of a socket insulator strip down to 3.5 holes (cut the middle of the fourth hole) with the cutting blade of a wire stripper. Scale down this segment to just 3 full holes and sand-down both sides if necessary. This helps to ensure that 3 full holes are available. Insert pin-wire units into the three holes of the micro strip until the crimped ends are flushed with the bottom edge of the insulator segment. The 10T assembly needs to be in one outer hole of the micro strip and not in the middle.
2. Modify bipolar stimulating electrodes (Figure 2).
   1. Give each rat one bipolar stimulating electrode. This electrode applies a small amount of electrical current via a stimulus isolator (see eyeblink system) as the shock US during ECC. The electrodes are acquired from a manufacturer in twisted form and are shielded.
   2. Untwist the two metal leads 2-3 times to make a V-shape (spread 5 mm) and then use the smooth high precision tweezer to straighten each "prong" as much as possible. Scrape off 1 mm of shielding from each prong's tip (all around) with a razor blade. Inspect each prong for irregularities and bends, and correct if necessary without scraping off additional shielding.
3. Remove manufacturing oil and foreign debris from the EMG headstages and bipolar electrodes by washing them in 95% ethyl alcohol. Allow them to dry then autoclave.

Figure 2: Electromyographic (EMG) recording electrodes and bipolar electrode. The finished EMG headstage (right, orange) is constructed from three male contact pins, two size 3T PTFE-coated wires, one size 10T PTFE-coated wire, and a modified micro strip. The three wires are approximately 5 cm each and are crimped to the contact pins. The finished bipolar electrode (right, white) is untwisted, re-straightened, and molded in a V-shape (5 mm split). Shielding is removed from the tips of the two prongs. Please click here to view a larger version of this figure.

4. Eyelid Surgery Procedure (Postnatal Day 90)

1. Preparation of materials (see Materials for detailed supply information), animals, surgical, and non-surgical areas:
   1. Autoclave surgical instruments and supplies - the number in parentheses ( ) indicates approximately how much is needed per adult rat or measurement: surgical instruments (1 set per 4 rats, as approved by the ECU IACUC), gauze pads (3-4), cotton-tipped swabs (5-6), 20 x 20 wrap (4 per/surgery session), porcelain crucible (1), stainless steel microspatula (1), 0.9% saline (1 wash bottle), Petri dish (1), and 0-80 stainless steel screws (3/rat).
   2. Other sterile supplies needed: surgical blade size 10 (1 per 4 rats), surgical drape (1 per rat), veterinary ophthalmic ointment, povidone-iodine (1 wash bottle), 26G x 3/8" hypodermic needle (2), nickel-plated pin vise with #55 drill bit, nickel-plated flathead jeweler's screwdriver (1.8-2 mm blade).
   3. Prepare a sterile holding area for all surgical tools and supplies using an approved research/medical grade disinfectant. Place materials on this space when the area is ready.
   4. Prepare a sterile surgical space using an approved research/medical grade disinfectant. This space contains a thermo-regulated water heating pad, stereotaxic apparatus fitted with a rat anesthesia mask, glass bead sterilizer, and isoflurane gas vaporizer (for anesthesia). The vaporizer has tubes to the anesthesia mask and to an induction chamber located on a separate space for non-sterile animal preparation procedures; each tube is controlled by its own valve.
   5. Scrub hands and arms thoroughly with antimicrobial soap. Without touching the inside, pre-open any items that have been packaged in sterilization pouches or wraps. Don sterile gloves using aseptic procedure and transfer all pertinent items to the surgical area. Set the drill bit at the proper depth in the pin vise (~ 2 mm) and prepare the screwdriver. Cover the sterile materials with a sterile wrap until the surgery is ready to begin.
   6. Prepare a non-sterile space with a weighing scale, electric fur trimmer, sterile 1 mL syringes with 26G x 3/8 in needles (1 per rat), surgery log, and buprenorphine (0.03 mg/mL concentration) administered 0.1 mL/100 grams.
      CAUTION: Buprenorphine is a DEA schedule III semisynthetic opioid, and must be locked and logged appropriately.
   7. Check the isoflurane vaporizer for appropriate gas level; add more gas if it is low. Leave the gas flow and mixture knobs off for now. Turn on the O₂ tank and check that there will be enough gas for all surgeries.
   8. Weigh each rat and record its body weight in the surgery log. Use a buprenorphine injection chart to determine the proper injection volume.
   9. Turn on the vaporizer valve to the induction chamber and leave the valve to the surgery mask off. Turn the mixture to 3 and flow rate to 3 (3% isoflurane with 100% O₂ as the carrier gas at a volume of 3 L/min).
   10. Place a rat in the induction chamber and allow it to reach proper anesthetic plane (slow breathing, lack of pedal reflex, lack of blink reflex). Remove the rat from the chamber once it has reached anesthetic plane.
   11. Shave its head using the fur trimmer, exposing a sufficient amount of skin for the incision site and the left eyelid. If necessary, place it back in the induction chamber to reach anesthetic plane again before resuming with the trimming.
   12. Disinfect the exposed skin by applying alternating rubs of isopropyl alcohol and povidone-iodine three times. If necessary, place it back in the induction chamber to reach anesthetic plane again, prior to transferring to the surgery table.
2. Surgery
   1. Turn on the vaporizer valve to the anesthesia mask and shut off the valve to the induction chamber. Adjust the mixture to 2.5 and flow rate to 2 (2.5% isoflurane with 100% O₂ as the carrier gas at a volume of 2 L/min). Transfer the rat to the stereotaxic apparatus and follow standard procedures in securing its head to the incisor bar and ear bars (see Geiger et al., 2008⁴⁹ for an example).
   2. Give the rat a pre-surgical injection of buprenorphine (SC) and carefully dispose of the needle (uncapped) in a sharps container. Place a sterile surgical drape on the rat, exposing just the surgical area and isolating it from the rest of the body.
   3. Don a surgical mask, surgical cap, surgical gown, goggles, and any other personal protective equipment required by the IACUC. Wash and scrub hands and arms thoroughly with antimicrobial soap. Don sterile gloves using sterile procedure.
   4. Apply a small amount of ophthalmic ointment to both eyes using a cotton-tipped swab to prevent them from drying.
   5. Proceed with the surgery when the rat exhibits proper anesthetic plane. Monitor the rat continuously throughout the surgery and check the isoflurane level periodically. Adjust the flow and mixture knobs accordingly during surgery based on the rat's response to the anesthesia.
   6. Make an anterior-posterior incision at the midline of the cranium with a scalpel blade. This incision should expose enough area in front of the eyes (i.e., exposing the frontal bone) and slightly behind the lambdoid suture.
   7. Scrape away the periosteum on top of the cranium carefully not to cause excessive bleeding. Wipe off excess connective tissue and blood with a cotton-tipped swab.
   8. Drill a hole using the drill bit with the aid of a pin vise, starting directly behind the coronal suture on one parietal bone. Remove any blood and bone debris with a cotton-tipped swab. Grasp a 0-80 screw by the threading with splinter forceps and fasten it to the hole with the jeweler's screwdriver; tighten-down the screw just enough without damaging cortical brain tissue (usually 3-4 full turns).
   9. Follow the same procedures described in 4.2.8 to fasten two more 0-80 screws to the skull, one directly behind the coronal suture of the opposite parietal bone and one directly anterior to the right lambdoid suture. It has been found that 3 screws are sufficient for anchoring the dental cement (see 4.2.18) to the cranium, as a fourth screw (anterior to the left lambdoid suture) would impede the placement of the bipolar electrode.
   10. Remove an EMG headstage from the Petri dish. Face the headstage so that the 10T wire is on the rat's right side. Bend the 10T wire upward, making it parallel with the bottom edge of the micro strip. Give it a slight bend to the right side so that the wire can come around the anterior and posterior 0-80 screws. Set it aside but close enough to reach.
   11. [Assuming one is right-handed] Place 3 in dressing forceps in the left hand and 4 in dressing forceps in the right hand. Grasp the upper skin at the incision site of its left eye with the 4 in dressing forceps and direct the 3 in dressing forceps towards the corner of this eye; grasp the skin at this corner. Maintain the grasp with the 3 in dressing forceps while releasing the 4 in dressing forceps.
   12. Take one 26G x 3/8 in needle and insert it through the corner of the eyelid; rotate the needle so that the beveled side is face-up. Use the micro dissecting forceps with platform to insert the middle 3T wire into the needle's hole. Push it through the hole a few centimeters without going down completely; this wire is the negative lead.
   13. Carry out the procedures described in 4.2.11 and 4.2.12 to insert the outside 3T wire into the middle portion of the eyelid; this wire is the positive lead.
   14. Grasp both needles with one hand while grasping the headstage with the other hand (or forceps). Pull the needles away from the eyelid while guiding the headstage in the same direction in one continuous movement. If necessary, rotate the headstage so that the 10T wire is towards the animal's right eye; do not allow the 3T wires to cross. Use fine tipped forceps to double-check that the positive lead is in the middle of the eyelid.
   15. Adjust the headstage so that it is centered between the eyes and is in an optimal position atop the frontal bone, anterior to the bilaterally-inserted screws. Hold the headstage with one hand and hold the 3 in dressing forceps with the other hand. Wrap the 10T wire around one or both screws on the 10T side.
   16. Place 3 in dressing forceps in one hand and 4 in dressing forceps in the other hand. Use the 3 in dressing forceps to grasp the skin at the incision site towards the left temple. Use the 4 in dressing forceps to create a small "pocket" by separating the skin (i.e., superficial fascia) from the temporalis muscle. Use the iris scissors to cut away more connective tissue, working in towards the corner of the left eye, to increase the size of this pocket. Be careful not to go in too deep or cut any blood vessels.
   17. Take a bipolar electrode and shape the wire leads so that they can be fitted along the curvature of the temporalis muscle, while the two prongs are situated (dorsal and ventral) posterior to the left eye (see video), and the bottom end of the bipolar electrode can be situated straight atop the cranium. Allow enough distance between the headstage and this electrode for the commutator cable plugs (Figure 3) to attach without impediment.
   18. Pour dental cement powder into the crucible - start with about 1/3 of the crucible, and then scale the amount as necessary. Add the liquid component until the consistency is watery, and mix these components with the spatula. When the cement's consistency becomes more viscous, apply it to the incision site - covering all edges of the skin with at least 1/8 in borders.
       1. Apply enough cement to fortify the headstage and bipolar electrode without preventing the rat from closing both of its eyes. Quickly wipe off any cement that drips onto unwanted areas of the rat's fur.
   19. Ensure that at least 4-5 threads of the bipolar electrode are left unsealed by cement so that the bipolar plug can be twisted on (Figure 3). Remove excess cement from the headstage - the top of the micro strip and the three contact pins must be clean. The cement hardens quickly and may easily prevent full coupling of any of these electrodes to the commutator cable plugs. Use splinter forceps to remove excess cement that has partly dried.
   20. Wait for the cement to harden; it becomes clear. Use the micro dissecting scissors to snip off excess length from the two 3T wires, leaving a few centimeters of working room.
   21. Grab the micro dissecting forceps with platform on the left hand while grabbing the micro dissecting forceps with fine points on the right hand. Grasp one 3T wire with the left forceps slightly in front of the eyelid. With the right forceps, push the eyelid up to expose more 3T wire and hold it there. Release the left forceps and use the left hand to grasp the headstage (ensure it is secure by now). Use the right forceps to strip off some PTFE shielding, which allows the wire to make contact with the orbicularis oculi muscle.
       NOTE: Take caution not to pull too hard on a 3T wire, as it may detach from the contact pin. If a wire detaches, then it will not be possible to replace the contact pin as the hardened cement will not release freely from the cranium without causing bone breakage.
   22. Perform the steps in 4.2.21 to strip off excess PTFE from the second 3T wire.
   23. Grab the two micro dissecting forceps as stated in 4.2.21 (with platform on left, fine points on right). Position the left forceps slightly in front of one 3T wire while leaving room for the right forceps to be positioned directly in front of the eyelid. Grasp the wire with both forceps.
       1. Create a "hook" by keeping the right forceps still and rotating the left forceps up and around towards the eyelid; do this in one motion. Carefully release the right forceps, and then the left forceps.
          NOTE: Hooking the electrodes on the eyelids help to minimize the possibility of them rubbing onto the cornea. This also minimizes the possibility for wire breakage during any point in the behavioral testing phase. While hooking does not fully prevent the wire from rubbing against the cornea for all animals, they typically show no signs of eye damage as reflected by their high blink amplitudes (measured by the digital oscilloscope in 5.1.7) and from daily post-surgical health observations (excessive tearing, partial eyelid closure, lack of mobility from eye damage). If a rat shows initial signs of discomfort, it is anesthetized with isoflurane (see 4.1.10) and its electrodes are re-examined and/or re-positioned. Ophthalmic ointment is applied to prevent the cornea from drying. The rat is observed daily for any post-procedural eye complications and should they persist, then seek veterinary care. Rats that exhibit severe eye complications which cannot be corrected with electrode re-positioning or with veterinary treatment, are humanely euthanized according to the standard operating procedures of the ECU IACUC.
   24. Perform the steps in 4.2.23 for the second 3T wire. Snip off excess wire using the micro dissecting scissors.
   25. Use one hand to release the rat from the ear bars while supporting its body with the other hand. Inspect its head for any blood or debris, clean up if necessary, and give it a post-operative injection of buprenorphine.
   26. Place the rat in a recovery bin that has a source of heat (such as a thermo-regulated water heating pad) to aid recovery. Mark in the surgery end time in the surgery log and any pertinent notes about the rat during surgery. Monitor the rat for proper respiration and when it exhibits sternal recumbency or moves about, it may be returned to its home cage.
   27. Re-sterilize the instruments (except micro dissecting forceps - as they may be damaged) using the bead sterilizer and ensure that the surgical area remains aseptic, and repeat aseptic procedures if multiple eyelid surgeries will be performed.

5. Trace Eyeblink Classical Conditioning Procedure

Figure 3: Modified operant conditioning box for eyeblink conditioning. Rats are freely-moving mammals, and therefore a rotating commutator is used for maintaining electrical signal contact from the EMG and bipolar plugs that are attached to the head. The commutator is attached to the arm of the stanchion, which is counter-weighted for alleviating pressure on the rat. A piezo tweeter (speaker) delivers a 2.8 kHz tone at 85 dB and these values are calibrated regularly. Acoustical foam assists with attenuating environmental noise. Please click here to view a larger version of this figure.

Figure 4: Eyeblink conditioning system. This custom-built system consists of an EMG Integrator unit that filters and amplifies incoming signals from the rats, a Stimulus Control unit that delivers various stimuli in addition to tones and shocks, a pre-amplifier for each operant box to increase EMG signal gain, and a stimulus isolator for each operant box; it provides varying shock levels (in mA). A digital oscilloscope (not part of the stock eyeblink system) is used for diagnostic purposes during habituation and acquisition. Please click here to view a larger version of this figure.

1. Begin habituation and acquisition training 5 days after surgery. Give each rat one day of handling and habituation to the training apparatus - consisting of a modified operant conditioning box that is housed in a larger sound-attenuated chamber (Figure 3) - and follow with 6 days of acquisition training. Carry out the habituation and acquisition sessions using a custom-built eyeblink system (Figure 4).
   1. Create a training log file and print it out to use for daily record keeping. This log should have the squad of rats pre-assigned pseudo-randomly (or other randomization method) to their training boxes.
   2. Power on the eyeblink system, computer, and run the eyeblink software. This version of the software has different modules (or applications) that run independently. Run the Animal module to create a *.RAT file that contains subject information (session number, animal ID, sex, weight, and box number). A single RAT file manages up to four rats, which comprises a squad of subjects. Enter the values accordingly and save the file. Create a separate RAT file for each squad of rats to be trained.
   3. Transfer the rats from their vivarium to the testing room and close the door. Handle each rat for 5 min.
   4. Transfer a rat to its designated box and connect the EMG and bipolar plugs from the commutator cable to the corresponding EMG headstage and bipolar electrode on its head.
      NOTE: On first exposure to the commutator cable, a rat may struggle and show signs of stress. Handle the rat carefully and allow breaks in between unsuccessful attachments to alleviate its stress.
   5. Use a commutator that rotates 360° while maintaining electrical signal delivery/reception as a rat moves freely within a box. It has 5 channels (3 for the EMG electrodes, 2 for the bipolar electrodes) leading into the EMG Integrator unit, which filters and amplifies the raw signal.
   6. Determine that the rat is moving freely, and that it is not hindered by the weight of the commutator cable assembly or stanchion arm. Adjust the counter weight at the back of the stanchion if necessary. Check that all plugs are secure then close the box door. Perform the same connection/check procedure for all rats.
   7. Use an oscilloscope to observe their EMG activity for abnormal signals (e.g., high frequency and/or high amplitude electrical activity) and a video surveillance system to check their status (e.g., motor activity, sensitivity to environment, signs of pain) if available. Note any problems that are observed in the training log.
   8. Allow rats to habituate to their chambers for 10 min, and then return them to their homecages.
   9. Clean and disinfect the operant boxes, sweep the floor, and clean the table(s). Carry out these procedures after every session.
2. Acquisition training occurs over 6 consecutive days (sessions).
   1. Day 1: Transfer a rat to its designated box and connect the EMG and bipolar plugs from the commutator cable to the corresponding EMG headstage and bipolar electrode on its head. Carry out the same procedures indicated in 5.1.6 and 5.1.7.
   2. Turn on the stimulus isolators and set the current to be delivered at 0.4 mA (4% of 10 mA as shown in video). The isolators deliver electrical current in alternating fashion (i.e., from trial to trial), between the dorsal and ventral prongs of the bipolar electrode. This helps reduce muscle fiber fatigue (at each muscle site) over repeated stimulus deliveries.
   3. Observe all eyeblink responses emitted and tolerances exhibited on each trial for all rats. After every second trial, increase the shock US intensity by 0.4 mA for each rat, until 1.6 mA is reached (on Trials 7 and 8). Double-check that all rats are responding normally in their boxes, that all amplification and gain settings are constant, and for proper electrical signals (i.e., low/little EMG noise) coming from them. Use a shock US intensity of 1.6 mA for all days of acquisition training.
      NOTE: If a rat exhibits pain to a given shock level (e.g., jumping high off the floor, climbing on walls, running around), the trial is paused and it will be returned to the previous mA setting (0.4 mA below) for two trials. Afterwards, the rat will receive the higher shock value that it did not tolerate again and if tolerance is shown, it will receive increasing values until 1.6 mA is reached. If it cannot tolerate any value, then the next lower value (in 0.4 increments) at which it tolerates will be used throughout training.
   4. Use an oscilloscope to observe their EMG activity for abnormal signals (e.g., high frequency and/or high amplitude electrical activity) and a video surveillance system to check their status (e.g., motor activity, sensitivity to environment, signs of pain).
      NOTE: Explanation of Trace ECC: A representative trial epoch with captured EMG waveforms is illustrated in Figure 1 and the trace ECC procedure is explained in detail in the video. The trace conditioning training file is set to deliver 100 trials (90 paired CS-US trials and 10 CS-only trials on every 10^th trial) with an average inter-trial interval of 30 sec. The entire training session lasts approximately 52 min (assuming no stoppages). All rats are naïve in Session 1 and will develop differential learning curves (as determined by the expression of CRs) as they progress through several days of training.
   5. Start the training: Run the Blink software module. When prompted, select the correct RAT file and the training file for "trace" eyeblink conditioning.
   6. Write down the start time and the experimenter's name in the training log. Monitor all activity at this point: proper EMG signals, good blink responses to the shock US (i.e., high/frequent URs), health status of the animals, and proper hardware function. Write any noteworthy remarks in the training log.
   7. Data collection is captured automatically by the software as RAW files (one per rat, per day). The RAW files contain EMG amplitude (in volts), frequency counts, latency, and areal data for startle responses (SRs), URs, and CRs (total and adaptive). Shock artifact obscures the first 100 ms of the UR period, therefore only the last 140 ms of the UR period contain UR data.
   8. At the end of training, the software will automatically close. Remove all rats from their training boxes and return them to the vivarium.
   9. Repeat steps 5.2.5 - 5.2.6 for all subsequent training days. Before starting the Blink module on a given day, update the session number within the RAT file. This allows for creation of new RAW files for all rats that day. Each rat should have 6 RAW files, assuming that none has been removed from the training.
3. Process the RAW files and create a file that has been filtered for statistical analysis.
   1. Run the Analysis module. Review each session for each rat on a trial-by-trial basis for problematic trials. This is carried out to screen for trials that may be compromised due to various reasons (e.g., excessive pre-CS baseline activity, excessive movement, electrode detachment, cabling/wire issues). The software has algorithms that assist the experimenter with detecting and removing "bad trials" from being part of the filtered data set.
   2. Import the filtered data into a spreadsheet program and perform pertinent statistical analyses.

结果

眨眼软件是能够提供一套大型和全面的数据对于许多类型的测量。为简洁起见，我们在此研究中，报告代表结果学习和性能包括自适应 CR 百分比、 自适应的 CR 振幅、 UR 百分比和 UR 振幅的措施。之所以选择自适应的 CR 期间，因为它代表着收购时宜眨眼反应过度重复训练，由于期间跟踪 ECC^50，，5152海马增强突触可塑性。UR 措施被选为澄清是否在新生儿酒精所致的学习赤字跟踪 ECC 的原因是联想学习中断或中断响应冲击我们-这可能表明动机或电机的差异，而不是学习处理群体之间的差异。为每个度量值的数据分析采用 2 （性别） x 3 （新生儿组） x 6 （会话） 混合不明，与会话作为重复测量因素。采用配对 t 检验的事后测试分析重要影响新生儿的治疗和显著的交互作用分析了使用简单效果测试。使用 0.05 最低 alpha 级别进行了所有的统计分析和图中的结果是平均 ± 镜

开始与自适应的 CR 百分比量值，方差分析表明显著的主效应，新生儿组， F(2，21) = 11.69， p < 0.001，但性没有显著的主效应 (p = 0.71) 或显著的交互作用，这些因素之间 (p = 0.20)。不出所料，自适应的 CR 百分比增加培训， F（5，105） 六届 = 81.15， p < 0.001 和新生儿群体之间的差异是依赖在某种程度上的会话， F（10，105） = 4.58， p < 0.001。没有其他显著的交互作用涉及会话因素。同样为自适应的 CR 振幅，那里又显著的主效应，新生儿组， F(2，21) = 22.32， p < 0.001，但性没有显著的主效应 (p = 0.21) 或显著的交互作用，这些因素之间 (p = 0.48)。CR 振幅也大大增加培训， F（5，105） 六届 = 59.27， p < 0.001 和新生儿群体之间的差异是依赖在某种程度上的会话， F（10，105） = 4.31， p < 0.001。总体来看，这两个 CR 措施有显着差异组和这些手段明显分隔在不同的会话的培训。若要确认哪些组差别很大，配对 t 检验的事后测试表明，酒精插管 (AI) 大鼠表现明显差于这两个 CR 措施比 unintubated-控制 (UC) 和深水插管 (SI) 大鼠 (p < 0.01 CR 百分比;p < CR 振幅为 0.001)，其中没有不与彼此不同 (p> 0.05)。重大的新生儿组 x 两个 CR 措施，确认 AI 大鼠显著更显著受损获取的 CRs 开始在会话 2 和贯彻会议 6 相比，大鼠 UC 和 SI 的会话交互执行简单的效果测试 (所有p< 0.05)，在六届会议期间不做从彼此不同。唯一的例外是自适应的 CR 振幅为寺大鼠才开始 AI 大鼠直到会话 3 相差很大。这些结果都显示在图 5A、 5B。

没有显著的差异，在 UR 措施由于性别、 新生儿组或与会话因素这些因素的交互。这些负面的结果表明，每个组能够发出对休克的眨眼反应我们同样，AI 大鼠模型观察学习赤字并不受激励或电机的闪烁 （图 6A、 6B） 的差别。

图 5: 跟踪采集条件响应 (平均 ± SEM)。早期的酒精暴露 （组 AI） 显著影响采集的自适应条件反应 (CR) 百分比 （A） 和 (B) 振幅。跟踪 ECC 是天生难以获得，因此这些措施是为所有的组-ECC 时滞相对较低，百分比可能会达到 80-85%的胎儿酒精紊乱症^21，53啮齿动物模型中。尽管如此，ECC 过程跟踪是征税对海马，这早期的脑发育过程中易受酒精影响的更多。* = p < 0.05，* * = p < 0.01，* * * = p < 0.001 之间 UC 和 AI 的大鼠;中括号提供了样本。请点击这里查看此图的大版本。

图 6: 无条件反应 （意思是 ± SEM） 采集。眨眼 （UR 百分比和 UR 振幅） 表现不组间差异显著。缺乏的差异表明在采集培训过程中使用的激波强度差异没有改变 AI 大鼠的动机或他们能够生产防御眨眼休克，相比两个对照组 （UC 和 SI） 的答复。中括号提供了样本。请点击这里查看此图的大版本。

讨论

在产后天 4 9 期间收到乙醇 （酒精） 的新生大鼠幼仔展出跟踪眨眼空调在成年后的损伤。这些发现支持酒精是致畸与持久对海马功能的有害影响的观点。总体而言，条件响应在跟踪过程中是大鼠暴露于酒精与两个对照组大鼠相比要低。联想学习障碍酒精暴露大鼠都不受激励或电机的差异 (即.，闪烁对美国冲击强度无差异)。

虽然跟踪 ECC 是阐明挑战诱导海马神经病理学会有用的诊断工具，这种方法的结果必须放入适当的上下文。第一，在本演示中的关键程序元素涉及酒精靶向的递送到发育中的大脑，允许录制的肌电活动和提供休克、 手术植入术的上述硬件，和随后的动物测试使用评估认知功能的兴趣学习范式的电极硬件制造漏洞已知窗口期间。在每个阶段的过程中，必须注意对啮齿动物的科目不造成不必要无意的伤害，并定期监测其健康体征。他们行为的结果向认知提供了"窗口"，准确地只是一个心理建构描述当他们的健康不会受到包括酒精加药，硬件缺陷或外科手术植入的实验误差。因此，在研究过程中的每个程序元素必须执行以声音的方式，以确保外推结果从 ECC 的同时，还可以到人体的研究结果。其次，ECC 范式提供了联想学习的性质，但必须小心不扩展使用这种方法的结果，大致所赋予它们到其他认知域-工作记忆、 短期长期记忆，等意识 — — 除非一已纳入这些域内 ECC 研究某些方面的实验设计。例如，此演示审查的追踪 ECC 学习，采集阶段，但未审查大鼠记忆保持他们完成培训后。内存因而是独立的心理过程，应该评估除了学习。按照设计，一个可能纳入记忆的保留间隔，以评估任何短期或长期的记忆能力。第三，有可能与动机、 经验性的和荷尔蒙因素促成的行为，同时工作的并行存储器系统⁵⁴的认识是必不可少的 （在 ECC) 相关性是的理解但揭示了什么是"好"或"差"关于学习的许多过程之一。最后，跟踪 ECC 不是纯粹海马依赖任务，其他的大脑区域可能介导 CR 的某些组件。因此，对不同的神经回路和/或在研究中，利用的刺激参数的类型之间相互作用的认识，必须考虑到当制作基于离散结果的影响。例如，小脑，也有助于跟踪 ECC，在那里它影响铬和铬时机，特别是三军情报局短时间的地形特点。痕量 ECC 在小脑受损的人用长的跟踪时间间隔 （1,000 女士），测试不受影响，但在那些接受较短的跟踪间隔 （400 毫秒）³⁴，影响。此外，背内侧前额叶皮层 （律） 的小鼠，前扣带回和内侧嗅沟地区为目标的双侧病变防止采集跟踪 CRs⁵⁵，而破坏的兔尾律产生类似结果⁴⁶。这些发现也突出考虑物种差异在小脑脑前额叶贡献阀杆驱动的联想学习，如跟踪 ECC 的重要性。而在 500 ms 跟踪 CRs 成人 PD 4 9 不利影响采集新生儿酒精暴露大鼠在这项研究和其他^47，56，这不是经验 300 ms 跟踪时间间隔，即使在较高剂量的酒精 （5 克/公斤）⁵⁷，暗示酒精染毒大鼠的微量损害依赖于跟踪间隔的挑战的新生儿酒精暴露大鼠的同一理由。

在此研究中，海马被强调是非常重要的调解 ECC，跟踪和新生儿酒精暴露受到挑战时展出相关神经损伤的缺陷跟踪 CRs 习得中可见一斑。但是，它必须告诫，小脑脑干电路，特别是间位核，是 ECC，包括采集、 表达式和地形特征的 CR，ECC 任务包括跟踪 ECC^{36，，4055，，5859}类型的许多方面的必要条件。事实上，这种神经电路与海马的驾驶中 ECC，如跟踪 ECC⁶⁰的高阶形式表达的 CRs 进行交互。是否早期大脑发育过程中的酒精暴露专门影响跟踪 ECC 中的海马功能尚不完全清楚。许多不同的大脑区域是易受早期酒精侮辱，包括律、 小脑、 海马^{18，19，，2347，61，62}，和很可能酒精扰乱跨许多 ECC 程序运作这些结构不同程度和不同，但功能上重要的差异。尽管关于从跟踪 ECC 研究结果解释的陷阱，跟踪 CRs 成功收购已被证明至少依靠完整的海马，动物病变研究^{42，44，63，，6465}所支持的。此过程因此仍然是非常有价值地演示之间发展酒精暴露于跟踪条件响应因为底层 it，神经回路要好得多的联系比其他海马相关的任务，如在 Morris 水迷宫，地方学习理解新物体再认，和上下文和跟踪担心空调。

ECC 作为行为法"测定"认知、 发展 neuroteratology 领域具有广泛的适用性。事实上，最近的调查结果，从我们的实验室支持发展中国家海马是酒精作用，可能通过不同的介入策略^18，47缓解对高度敏感的概念。这里的关键的好处是与酒精所致的微量 ECC 学习赤字更好地了解，他们可能是预测中海马基于功能联想学习-以外的其他问题的特别是那些已知的由同一海马神经介导。

应用微量 ECC 和及其其他变体 （例如，延迟、 逆转、 歧视、 复合） 澄清的神经生物学机制和神经系统参与的联想学习，还可以扩展超出了胎儿酒精研究领域。例如，这种范式在人类病例及动物模型，精神疾病如精神分裂症^66，67、 神经退行性疾病，如老年痴呆症^68，69，以及药物的滥用^70，，7172关注。它作为一种研究方法来评估认知功能和功能障碍的好处也由此明显横跨许多心理和生物医学的学科，包括神经科学。

材料

Name	Company	Catalog Number	Comments
Neonatal Alcohol Exposure
190 Proof Ethyl Alcohol (USP)	Pharmco-AAPER	225-36000 [ECU Medical Storeroom]	Can be substituted; should be USP; avoid using 200 proof ethyl alcohol
Container/Basket for Pups	Any
Corn Oil	Any		Food grade
Heated Water Therapy Pump w/ Pads	Gaymar	TP-500	To keep pups warm; can be substituted
Hypodermic Needles 22G x 1 in, Sterile	Any
Hypodermic Needles 30G x 1/2 in, Sterile	Any
Isopropyl Alcohol 70%	EMD Millipore	PX1840-4 [Fisher Scientific]	Can be substituted; reagent grade www.fishersci.com
Long-Evans Rats (Female and Male Breeders)	Charles River Laboratories	N/A [ECU Dept. of Comparative Medicine]	Age and weight need to be specified; pricing varies by these factors www.criver.com
Micro Dissecting Scissors, 3.5 in, 23 mm Blades	Biomedical Research Instruments	11-2200	For cutting PE tubing brisurgical.com
Polyethylene 10 Tubing (0.011 in. I.D.; 0.024 in. O.D.)	BD Diagnostic Systems	22-204008 [Fisher Scientific]	Can be substituted www.fishersci.com
Polyethylene 50 Tubing (0.023 in. I.D.; 0.038 in. O.D.)	BD Diagnostic Systems	22270835 [Fisher Scientific]	Can be substituted www.fishersci.com
Regulated water heater or baby milk bottle warmer	Any		Optional; helps with warming up cold milk solutions
Tuberculin Syringes, Sterile, 1.0 ml	Any
Tuberculin Syringes, Sterile, 10 ml	Any		Can be used to draw out ethyl alcohol or use appropriate size micropipet
Weigh Scale	Any		Should have good resolution (in gram units)
Name	Company	Catalog Number	Comments
EMG Headstage Fabrication and Bipolar Electrode Modification
Bipolar Electrode, 2 Channel SS Twisted	Plastics One, Inc.	MS303/2-B/SPC  ELECT SS  2C TW .008"	Must specify custom length of 20 mm below pedestal www.plastics1.com
Centi-Loc Strip Socket Insulator (aka, Micro Strip)	ITT Cannon / ITT Interconnect Solutions	CTA4-IS-60* or CTA4-1S-60*	*Depends on vendor; see www.onlinecomponents.com or www.avnetexpress.avnet.com
Dental Pliers, Serrated	CMF Medicon	390.20.05	Can be substituted; use to crimp wires to male contact pins www.medicon.de
Micro Dissecting Scissors, 3.5 in, 23 mm Blades	Biomedical Research Instruments	11-2200	Only use to cut 3T wires; cutting 10T wires will damage the blade - use the blade of the wire stripper instead brisurgical.com
PTFE-Coated Stainless Steel Wire, 10T (Bare Diameter .010 in)	Sigmund Cohn-Medwire	316SS10T	www.sigmundcohn.com
PTFE-Coated Stainless Steel Wire, 3T (Bare Diameter 0.003 in)	Sigmund Cohn-Medwire	316SS3T	www.sigmundcohn.com
Razor Blade	Any		To strip 1 mm from prongs of bipolar electrode
Relia-Tac Socket Contact Pin, Male	Cooper Interconnect	220-P02-100	See Allied Electronics Cat # 70144761 www.alliedelec.com
Tweezers, High Precision, Serrated, 4 3/4 in	Electron Microscopy Sciences	78314-00D	To grasp 10T wire firmly while stripping PTFE with smooth tweezers www.emsdiasum.com
Tweezers, High Precision, Smooth, 4 3/4 in	Electron Microscopy Sciences	78313-00B	www.emsdiasum.com
Tweezers, Ultra Fine Tips, 4 3/4 in	Electron Microscopy Sciences	78510-0	To strip 1 mm of PTFE from one end of 3T wire; grasp shielded portion with smooth tweezers www.emsdiasum.com
Wire Stripper, 16-26 AWG	Any		Use the blade end to cut micro strips
Name	Company	Catalog Number	Comments
Eyelid Surgery
Surgical Instruments (High Quality Stainless Steel)
2 x Dressing Forceps, 4 in Serrated	Biomedical Research Instruments	30-1205	Can be substituted; extra forceps for grasping electrodes/screws outside of surgery tray brisurgical.com
Dressing Forceps, 3 in Serrated	Biomedical Research Instruments	30-1200	Can be substituted brisurgical.com
Instrument Tray	Biomedical Research Instruments	24-1355	Can be substituted brisurgical.com
Knife Handle No. 3, 5 in	Biomedical Research Instruments	26-1000	Can be substituted brisurgical.com
Micro Dissecting Forceps, 3.5 in, Fine Points	Biomedical Research Instruments	10-1630	Can be substituted brisurgical.com
Micro Dissecting Forceps, 3.5 in, Smooth Platform (0.3 x 5 mm)	Biomedical Research Instruments	10-1720	brisurgical.com
Micro Dissecting Scissors, 3.5 in, Extremely Delicate, 15 mm Blades	Biomedical Research Instruments	11-2000	Can be substituted brisurgical.com
Plain Splinter Forceps, 3.5 in	Biomedical Research Instruments	30-1600	Can be substituted brisurgical.com
#10 Stainless Steel Surgical Blade for #3 Handle, Sterile	Any		Can be substituted
0-80 x 0.125 in Stainless Steel Screws	Plastics One, Inc.	0-80 x 0.125	Can be substituted www.plastics1.com
Alcohol Prep Pads, Sterile	Fisher Scientific	22-363-750 [Fisher Scientific	Can be substituted www.fishersci.com
Betadine Povidone-Iodine	Purdue Frederick Co.	6761815101 [Fisher Scientific]	Can be substituted www.fishersci.com
Betadine Povidone-Iodine Prep Pads	Moore Medical	19-898-946 [Fisher Scientific]	Can be substituted www.fishersci.com
Cotton-Tipped Swabs, Autoclavable	Any		Typically 7.6 cm or 15.2 cm length
Drill Bit for Pin Vise, #55 (0.052 in)	Any		Metal should resist rusting and corrosion
Gauze Pads, 2 in x 2 in	Fisher Scientific	22-362-178 [Fisher Scientific]	Can be substituted www.fishersci.com
General Purpose Latex/Nitrile/Vinyl Gloves	Any
Glass Bead Sterilizer	Any		Sterilize instruments between surgeries
Heated Water Therapy Pump w/ Pads x 2	Gaymar	TP-500	Can be substituted; separate pumps are recommended - 1 for surgery, 1 for recovery
Hypodermic Needles 26G x 3/8 in, Sterile	Any
Isoflurane	Vedco	NDC 50989-150-12	Manfacturer can be substituted; veterinary approval may be required
Isoflurane Vaporizer System, Tabletop, Non-Rebreathing	Parkland Scientific	V3000PK	Can be substituted www.parklandscientific.com
Jewelers Screwdriver w/ 1.8-2 mm Blade	Any		Metal should resist rusting and corrosion
Ortho-Jet BCA Package (Dental Cement)	Lang Dental	B1334	Contains powder (1 lb) and liquid www.langdental.com
Oxygen Tank with Pressure Regulator, Large			Local supplier
Porcelain Crucible, High-Form, Glazed, 10 ml	CoorsTek, Inc.	07-965C [Fisher Scientific]	Can be substituted with Fisher FB-965-I Wide-Form Crucible www.fishersci.com
Puralube Veterinary Ophthalmic Ointment, Sterile	Henry Schein Company	NC0144682 [Fisher Scientific]	Can be substituted www.fishersci.com
Quatricide PV-15	Pharmacal	PV-15	Antimicrobial disinfectant; can be substituted www.pharmacal.com
Rat Gas Anesthesia Masks for Stereotaxic Surgery	Stoelting Company	51610	www.stoeltingco.com
Rat Stereotaxic Apparatus w/ Ear Bars (45 Degree)	Any		45 degree bars are recommended to prevent damaging eardrums
Roboz Surgical Instrument Milk	Roboz Surgical	NC9358575 [Fisher Scientific]	Can be substituted; for lubricating instruments during autoclaving www.fishersci.com
Rodent Hair Trimmer	Any
Sodium Chloride	Fisher Scientific	S641-500 [Fisher Scientific]	To make 0.9% saline; reagent grade; USP www.fishersci.com
Stainless Steel Microspatula (Blade: 0.75 L x 0.18 in. W)	Fisher Scientific	21-401-15 [Fisher Scientific]	Can be substituted www.fishersci.com
Starrett Pin Vise, 0.000 in - 0.055 in	Any		Nickel-plated or equivalent recommended to resist rusting and corrosion
Sterile Surgical Gloves	Any
Sterilization Wraps, 20 in x 20 in, Autoclavable	Propper Manufacturing	11-890-8C [Fisher Scientific]	Useful for wrapping autoclavable supplies and on sterile field during surgery www.fishersci.com
Surgical Drape, Sterile/Autoclavable	Any		May need to cut to size for rats
Surgical Gown*	Any		*If required by IACUC
Surgical Mask	Any
Tuberculin Syringes, Sterile, 1.0 ml	Any
Weigh Scale	Any		Should have good resolution (in gram units)
Name	Company	Catalog Number	Comments
Eyeblink System and Components (assuming 4-rodent system)
5 Channel Commutator x 4	Plastics One, Inc.	SL2 + 3C	www.plastics1.com
Bipolar Electrode Cable, Dual 305 x 4	Plastics One, Inc.	305-305 80CM TT2 (C)	Provides plug end to bipolar electrode on rat and to commutator; must be modified www.plastics1.com
Cable, 5 Channel, Shielded, 26 AWG x 4	Any		To fabricate commutator cable; this must be made from scratch
Chamber for Operant Test Box (Inside: 24 H x 23 W x 14 D in) x 4	Med-Associates		Can be substituted; inner dimensions should fit operant test box comfortably, with room for acoustical foam; fit with fan - 55-60 dB www.med-associates.com
Eyeblink System and Software	JSA Designs	N/A	Proprietary and customized for research lab
Heat Shrink Tubing (3/16 in, 1/4 in, 3/8 in, 1/2 in Diameters)	Any		To protect modified commutator cable soldered ends and splices
Melamine Triple Peak Acoustical Foam w/Black Hypalon (24 x 48 in)	McMaster-Carr	9162T5	Can be substituted; cut to fit 4 housing chambers www.mcmaster.com
Operant Test Box (Exterior 12.5 L x 10 W x 13.5 in H), Complete x 4	Med-Associates	ENV-007 Custom Package	With stainless steel grid floor and custom top (3 in hole in center for commutator cable) www.med-associates.com
Oscilloscope (Optional)	Any		Recommended minimum specs: 200 MHz analog bandwidth, 1 GS/s real-time sampling, 4 channels; see www.picotech.com /td>
Piezo Tweeters (Speakers) x 4 (7 x 3 in)	MCM Electronics	53-805	Must match frequency range specifications for eyeblink system (2500 Hz - 25 KHz) www.mcmelectronics.com
Soldering Station, Solder, Flux, Tinner	Any		For soldering 26 AWG cables to female sockets (that fit male relia-tac contact pins) and bipolar plugs
Stimulus Isolators x 4	WPI International	A365	These units run on 16-9V alkaline batteries; a suitable rechargeable version (A365R) is available www.wpiinc.com
Tripolar Electrode Cable for SL3C Commutator x 4	Plastics One, Inc.	335-335 80cm TT3 C	Provides plug end to EMG headstage on rat and to commutator; must be modified www.plastics1.com
USB LED Lights x 4	Any		USB-based lights do not cause electrical "noise" with the EMG signals from the rats www.plastics1.com
Webcams x 4, Surveillance Software	Any
PC Computer Running MS Windows OS	Any