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Neurociência

胎児アルコール スペクトルの無秩序のモデルラットにおける海馬機能障害を評価するためにトレース瞬目反射条件づけの使用

Published: August 05, 2017
doi:
10.3791/55350
, , , , ,
1Department of Psychology,East Carolina University, 2Multidisciplinary Studies Program in Neuroscience,East Carolina University

Summary

トレース瞬目反射条件づけ (ECC) を用いて評価したラット海馬依存学習投与早期新生児の脳の発達の中にアルコールの高濃度 (11.9 %v/v)。一般的には、ECC プロシージャは、多くの心理的な医学の設定の間で脳機能障害を検出するため音の診断ツールです。

Abstract

新生児ラット生後 4-9 中に比較的高い濃度エチルアルコール (11.9 %v/v) の胎児の脳が急速な組織変化を起こしのようであると時間加速人間の 3 番目の妊娠中に発生した脳の変化。胎児アルコール スペクトルの無秩序 (FASDs) のこのモデルは、量およびいくつかの妊娠中のアルコールの母親で発生するどんちゃん騒ぎの飲むのパターンを模倣した重症の脳損傷を生成します。我々 はトレース瞬目反射条件づけ (ECC)、アルコール露出アダルト子孫に通常見られる長期の海馬の機能不全を評価するために、連合学習の高次変形の使用をについて説明します。90 日齢、齧歯動物は手術記録と準備ができていたと筋電図 (EMG) を測定した刺激的な電極左目のまぶたの筋肉から活動を点滅し、左目の後方は穏やかなショックをそれぞれ配信。5 日間の回復期間後彼らはトレース アルコール曝露し、ラットを制御の連合学習の違いを決定する ECC の 6 セッションを受けた。トレース ECC は、簡単に変更できる同じ機器やソフトウェアを使用して異なる神経系を評価することができますので、多くの可能な ECC 手順の 1 つです。一般的には、ECC 手順別の脳の仕組みと脳を侮辱したさまざまな条件で神経病理学を検出するため、診断ツールとして使用できます。

Introduction

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 severe3,4. 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 learning9,10 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 4-912,13, a period when the brain undergoes rapid synapse and dendritic contact formation, comparable to human fetal week 24 and extending into the 3rd trimester14,15,16,17. This particular model induces significant loss of hippocampal neurons18,19 and neurons in many other brain regions such as the cerebellum12,13,14,15,16,17,18,19,20,21,22,23, accompanied by severe impairments in cognitive functions spanning different domains21,24,25. 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 learning26,27 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 development28,29,30,31. Furthermore, the fundamental neural circuits that mediate associative learning in this paradigm are supported by experimental and neuropsychological reports in these same species26,32,33,34,35,36,37.

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-circuitry38,39,40, whereas trace ECC is a higher-order procedure that requires additional neural engagement from the hippocampus41,42,43,44 and other cortical structures45,46. 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
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 exposure18,47. 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.

Protocol

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 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. 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. Use micro dissecting scissors to cut a 3 in piece of PE-50 tubing and a 6 in piece of PE-10 tubing. Carefully attach the PE-50 tubing to a sterile 22 G x 1 in hypodermic needle. 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. 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. Store the complete intragastric intubation tube in fresh 70% isopropyl alcohol to keep it sterile. 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. 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. Shake this bottle well and label it accordingly (i.e., 11.9% v/v Ethyl Alcohol in Milk). 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. Store both milk bottles in the refrigerator. 2. Neonatal Alcohol Exposure (Postnatal Days 4-9) 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. 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. 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. 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. Place the 11.9% v/v Ethyl Alcohol in Milk solution in a warm water bath and shake it well. Intragastric intubation procedure: Flush out the intragastric intubation tube that was stored in 70% isopropyl alcohol well with warm water. Extract the correct amount of solution using a sterile 1 mL syringe. Dip the tip of the intubation tube (PE-10 end) in fresh corn oil (this facilitates insertion). 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. 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. 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). 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). 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). 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. 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. 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 Fabricate one electromyographic (EMG) "headstage" for each rat (Figure 2). The headstage allows for recording of eyeblink responses via the eyeblink system (Figure 4). 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). 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. 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. 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. Perform tug tests on all wires to ensure that they are secured to the pins. Take caution not to bend or damage the wires. 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. Modify bipolar stimulating electrodes (Figure 2). 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. 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. 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) Preparation of materials (see Materials for detailed supply information), animals, surgical, and non-surgical areas: 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). 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). 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. 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. 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. 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. 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. Weigh each rat and record its body weight in the surgery log. Use a buprenorphine injection chart to determine the proper injection volume. 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). 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. 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. 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. Surgery 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). 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. 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. Apply a small amount of ophthalmic ointment to both eyes using a cotton-tipped swab to prevent them from drying. 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. 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. 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. 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). 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. 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. [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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. 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. Perform the steps in 4.2.21 to strip off excess PTFE from the second 3T wire. 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. 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. Perform the steps in 4.2.23 for the second 3T wire. Snip off excess wire using the micro dissecting scissors. 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. 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. 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. 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). 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. 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. Transfer the rats from their vivarium to the testing room and close the door. Handle each rat for 5 min. 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. 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. 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. 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. Allow rats to habituate to their chambers for 10 min, and then return them to their homecages. Clean and disinfect the operant boxes, sweep the floor, and clean the table(s). Carry out these procedures after every session. Acquisition training occurs over 6 consecutive days (sessions). 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. 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. 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. 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 10th 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. Start the training: Run the Blink software module. When prompted, select the correct RAT file and the training file for "trace" eyeblink conditioning. 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. 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. At the end of training, the software will automatically close. Remove all rats from their training boxes and return them to the vivarium. 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. Process the RAW files and create a file that has been filtered for statistical analysis. 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. Import the filtered data into a spreadsheet program and perform pertinent statistical analyses.

Representative Results

瞬目ソフトウェア測定の多くの種類の大規模な包括的なデータのセットを提供することが可能です。簡潔にするため、本研究で報告代表結果学習とパフォーマンスの適応の CR 割合、適応 CR 振幅、UR 割合 UR 振幅などの措置。適応 CR 期間は、ECC50,51,52トレース中の海馬のシナプス可塑性の強化の結果として、繰り返しのトレーニング、時宜を得た瞬目反応の獲得を表すために選ばれました。UR 対策で新生児のアルコール性の学習の欠損トレース ECC 治療群学習相違点ではなく、動機づけやモーターの違いを示す可能性があります – 私たち連合学習の混乱やショックへの対応に混乱が原因だったかどうかを解明するため選ばれました。各メジャーのデータは、混合性、繰り返し対策要因としてセッション 2 (セックス) x 3 (新生児グループ) x 6 (セッション) を使用して分析しました。新生児の治療のため有意な主効果テューキー事後テストを使用して分析し、重要な相互作用を単純な効果テストを使用して行った。0.05 の最小アルファ レベルを使用してすべての統計解析を実施し、グラフの結果が平均 ± SEM. 分散適応の CR 割合測定に始まり、新生児グループ、 F(2,21) の有意な主効果を示される = 11.69、 p < 0.001、しかしセックスのない有意な主効果 (p = 0.71) またはこれらの要因間の重要な相互作用 (p = 0.20)。予想通り、適応の CR 割合増加トレーニング、 F(5, 105) の六つのセッション = 81.15、 p < 0.001 と新生児グループ間の違いはF(10, 105) セッションのいくつかのレベルに依存していた 4.58、 p = < 0.001。セッション因子を含む他の重要な相互作用はありませんでした。同様に適応の CR 振幅があった再びF(2,21) 新生児グループの有意な主効果 22.32、 p = < 0.001、しかしセックスのない有意な主効果 (p = 0.21) またはこれらの要因間の重要な相互作用 (p = 0.48)。CR 振幅も大幅にトレーニング、 F(5, 105) の六つのセッションを介して増加 59.27、 p = < 0.001 と新生児グループ間の違いはF(10, 105) セッションのいくつかのレベルに依存していた 4.31、 p = < 0.001。全体的にみて、両方 CR 対策グループ手段および訓練の別のセッションで大きく区切られたこれらの手段の間で有意差を示した。グループが大幅に異なることを確認、するテューキー事後テストに unintubated 制御 (UC) と偽挿管 (SI) ラットよりも両方の CR の措置に対して、アルコール挿管 (AI) ラット実行大幅に悪化を示した (p < 0.01 の CR 割合;p < CR 振幅の 0.001)、互いを認められなかった (p> 0.05)。単純な効果のテストは、x セッションの相互作用両方 CR の措置については、AI ラットが UC と SI の両方のラットと比較して始まるセッション 2、セッション 6 を乗せて CRs の買収により著しく損なわれたことを確認重要な新生児グループに対して実行される (すべてp< 0.05)、互いから六つのセッションを通して異ならなかった。唯一の例外は、セッション 3 まで AI ラットと大幅に異なる SI ラットが始まらなかったため適応 CR 振幅をだった。これらの結果は、図 5 a、 5 bのとおりです。 セックス、新生児グループ、またはセッション因子のこれらの要因の相互作用のため UR 対策に有意差はありませんでした。これらの陰性所見では、各グループは私たち同様に、瞬目ショック レスポンスを生成することができたし、(図 6 a、 6 b) 点滅の動機付けや運動の違いによって影響を受ける AI ラットで観察学習の欠損がなかったことを示されています。 図 5: トレースの取得エアコン応答 (平均 ± SEM).初期のアルコール暴露 (AI グループ) には、適応条件反応 (CR) の割合 (A) と (B) の振幅による著しい影響を受けます。トレース ECC は本質的に取得することは困難、したがって対策は ECC の遅延 – すべてのグループの比較的低い、パーセンテージは FASD21,53の齧歯動物モデルの 80-85% に達する可能性があります。それにもかかわらず、トレース ECC プロシージャは、初期の脳の発達中にはアルコールの影響を受け、海馬で課税の詳細です。* = p < 0.05 * * p = < 0.01、* * * p = < UC と AI のラット間 0.001サンプル サイズは、かっこ内に提供されます。この図の拡大版を表示するのにはここをクリックしてください。 図 6: 無条件反応 (± SEM を意味する) の取得します。瞬目パフォーマンス (UR 割合と UR 振幅) は、グループ間に有意差でした。違いの欠如は、その習得訓練中に使用される衝撃強度に差動 AI ラットの動機は影響しなかったか守備能力点滅 (UC と SI) 両方の制御グループと比較して、衝撃への応答を示します。サンプル サイズは、かっこ内に提供されます。この図の拡大版を表示するのにはここをクリックしてください。

Discussion

生後 4-9 中にエチルアルコールを受信したラット新生仔は、成人の海馬障害を出展しました。これらの調査結果は、アルコールが海馬の機能に有害な影響を永続的の催奇形物質であるという考えをサポートします。両方のコントロールのグループのラットと比較してアルコールにさらされたラットのトレースの手順で全体的に、調節された応答が低かった。アルコールに曝露されたラットの学習障害が動機またはモーターの違いの影響を受けなかった (すなわち。、点滅米国の衝撃強度に違いはありません)。

トレース ECC 解明課題による海馬神経病理学の有用な診断ツールですが、このメソッドの結果は、適切なコンテキストに置かれなければなりません。最初に、このデモでキーの手続き型の要素には、発展途上の脳筋活動電位の記録ができ、衝撃、前述のハードウェア、および関心の認知機能を評価する学習パラダイムを使用してその後動物テストの外科的移植を提供する電極ハードウェアの作製に脆弱性の既知のウィンドウの中にアルコールのターゲットを絞った配信が関与しています。プロセスの各段階で注意する必要がある齧歯動物の被験者に不要な/意図しない害を発生しないようにし、定期的に自分の健康の兆候を監視します。彼らの行動の結果は、認知の「ウィンドウ」は、正確には心理的な構造説明とき自分の健康を含むアルコール投与、ハードウェアの欠陥、または外科的移植実験の誤差によって妥協されません。したがって、研究プロセスの各手順の要素は人間の所見に ECC からの結果を推定できることを保障するために健全な方法で実行されなければなりません。第二に、ECC パラダイム連合学習の性質に洞察力を提供しますが、1 つが実験的なデザインで ECC 研究内のこれらのドメインのいくつかのファセットを組み込んでいる限りは、このアプローチを使用して結果を拡張し、広くそれらを他の認知ドメイン – 作業メモリ、短期/長期記憶、意識などに帰するに注意が必要があります。たとえば、このデモはトレース ECC 学習の獲得の段階を検討したが、彼らは訓練を完了した後、ラットにおける記憶保持を調査しませんでした。メモリは、学習だけでなく評価される独立した心理的プロセスです。仕様では、1 つはいずれかの短期または長期の記憶能力を評価するためにメモリの保存期間を組み込むことができます。第三に、並列メモリ システム54の動作に寄与する要因を動機づけ、経験的なおよびホルモンと一緒に同時に動作可能性がありますがあることの認識は、(ECC) の間のアソシエティビティが理解が、その正体は「良い」または「悪い」学習について多くのプロセスの 1 つに不可欠です。最後に、トレース ECC は他の頭脳領域は、CR のいくつかのコンポーネントを仲介することがありますので、純粋な海馬依存タスクではありません。したがって、異なる神経回路および/または研究で使用されている刺激パラメーターの型間の相互作用を理解する必要があります考慮する離散の結果に基づいて影響を作るとき。小脳などもトレース ECC、CR とタイミング、特に ISI は短い期間で CR の地形学的特徴に影響を及ぼす場所に貢献します。トレース ECC は長いトレース間隔 (1,000 ミリ秒) でテストが小脳損傷に人間は影響を受けませんが、短いトレース間隔 (400 ms)34を受給している人の影響を受けます。さらに、マウスの前部帯状回と内側の agranular 地域をターゲットに、背内側前頭前野 (mPFC) の両側性病変は、ウサギの尾 mPFC の破壊生成と同様の結果46CRs55トレースの取得を防ぐ。これらの調査結果はまた小脳脳前頭貢献に種差幹トレース ECC などの駆動の連合学習を考慮の重要性を強調表示します。大人用 500 ms トレース CRs の PD 4-9 悪影響を取得中に新生児のアルコール暴露は47,56本研究でラット、これは 300 ms トレース間隔が発生するアルコール (5 g/kg)57, アルコール曝露ラットにおけるトレース障害がトレース間隔の時間に依存することを示唆の比較的高用量で挑戦されたときにも新生児のアルコールに曝露されたラットに同じケースではありません。

本研究では海馬はトレース CRs の習得における障害を反映して神経に関連する損傷を展示トレース、ECC を媒介と新生児アルコール露出によって挑戦されたとき極めて重要であることを強調しました。ただし、ECC、買収、式、トレース ECC36,40,55,58,59を含む ECC タスクの種類に応じて、CR の地形などの多くの面は特に規核、小脳脳幹回路が欠かせませんそれ警告する必要があります。確かに、この神経回路はトレース ECC60など、ECC の上位フォーム時に CRs の式を運転の海馬と対話します。初期の脳の発達中のアルコール暴露はトレース ECC で海馬の機能、特に効果かどうかは、全く明らかではないです。多くの異なる脳の領域は、mPFC、小脳、海馬18,19,23,47,,6162など初期のアルコール侮辱を受けます、アルコールの妨害が多く ECC プロシージャ間程度と変えることで、機能的に重要な相違点がこれらの構造体の機能は非常にそうです。トレース ECC 研究の結果の解釈に関する落とし穴にもかかわらず動物病変研究42,44,63,64,65でサポートされている、少なくともそのまま海馬に依存するトレース CRs の買収に成功を示されています。この手順はこうして残る新規オブジェクト認識、モリス水迷路学習の場所など、他の海馬依存タスクのよりも理解したトレース条件、それの基礎となる神経回路ははるかに良いために、応答する発達アルコール露出間のリンクを示すために非常に貴重なアプローチおよびコンテキストとトレースの恐怖します。

「測定」認知行動法として ECC 発達 neuroteratology の分野で広範な適用性があります。確かに、当研究室の最近の知見は、発展途上の海馬はアルコールの効果は、さまざまな介入戦略18,47によって緩和されるかもしれませんに機密性の高い概念をサポートします。ここでの主な利点かもしれないということですアルコール誘起トレース ECC 学習の欠損のよりよい理解、彼ら連想学習 – 外海馬機能のほかの問題の予測同じ海馬 neurocircuitry によって媒介される知られている特にそれら。

アプリケーション トレース ECC およびその他の変種 (例えば遅延、反転、差別、化合物) 神経生物学メカニズムと連合学習に関係する神経システムを解明するは、胎児性アルコール研究分野を超えて拡張できます。たとえば、このパラダイムは、症例と6968,アルツハイマー病など神経変性疾患、統合失調症66,67などの精神疾患の動物モデル、薬物乱用70,,7172で注目を受けています。したがっています神経科学を含む多くの心理的な生物医学分野の neurocognitive 機能と機能障害を評価するための研究方法とその利点を明らかに。

Declarações

The authors have nothing to disclose.

Acknowledgements

この作品は、アルコール飲料医療研究財団 (ABMRF) から TDT に助成金によって支えられました。

Materials

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
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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
DOWNLOAD MATERIALS LIST

Referências

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Keywords: Trace Eyeblink Classical ConditioningHippocampal DysfunctionFetal Alcohol Spectrum DisordersNeonatal Rat PupsAlcohol ExposureThird TrimesterHippocampusAssociative LearningBinge Exposure ParadigmAlcohol SolutionPE-10 TubeEsophagusGastroesophageal Sphincter
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Tran, T. D., Amin, A., Jones, K. G., Sheffer, E. M., Ortega, L., Dolman, K. The Use of Trace Eyeblink Classical Conditioning to Assess Hippocampal Dysfunction in a Rat Model of Fetal Alcohol Spectrum Disorders. J. Vis. Exp. (126), e55350, doi:10.3791/55350 (2017).
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