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Sep 2018

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Inducing Alcohol Dependence in Rats Using Chronic Intermittent Exposure to Alcohol Vapor
长期间歇性暴露于酒精蒸汽以诱导大鼠酒精依赖   

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Abstract

Alcohol use disorder (AUD) is a significant public health and economic burden and is often characterized by repeated bouts of alcohol intoxication and withdrawal. Virtually all organ systems are impacted by chronic alcohol exposure. These effects can be investigated using the rat as a model organism; however, rats typically will not self-administer alcohol to levels necessary to achieve physiological and behavioral aspects of dependence. The protocol described herein can be utilized to induce alcohol dependence in rats by administering alcohol vapor to the homecage for an extended period of time. This method allows the researcher to control the level, duration, and pattern of intoxication, and it reliably induces physiological and behavioral aspects of alcohol dependence, allowing for the study of biology and behavior with relevance for AUD in humans.

Keywords: Alcohol dependence (酒精依赖), Alcohol use disorder (酒精使用障碍), Vapor (蒸汽), Withdrawal (脱瘾), Intoxication (中毒), Blood alcohol level (血液酒精水平), Rat (大鼠)

Background

Alcohol use disorder (AUD) affects over 15 million Americans (SAMHSA, 2015), costing the US ~$249 billion annually (Sacks et al., 2015). The progressive development of AUD is characterized by repeated periods of alcohol intoxication and withdrawal, which initiates a series of neuroadaptations and behavioral changes. Individuals with AUD drink excessive quantities of alcohol, partly because alcohol has both positive (i.e., drinking alcohol for its rewarding effects) and negative (i.e., drinking alcohol to remove or avoid aversive feelings) reinforcing effects in those individuals (Koob, 2003). Alcohol-dependent rats withdrawn from alcohol exhibit many of the same behaviors exhibited by alcohol-dependent humans, including increases in alcohol drinking (e.g., Roberts et al., 1996), motivation to obtain alcohol (e.g., Walker and Koob, 2007), anxiety-like behavior (e.g., Kallupi et al., 2014) and nociception (e.g., Egli et al., 2012; Avegno et al., 2018). These behaviors suggest that rat models of alcohol dependence are a useful tool for investigating the neurobiological underpinnings of alcohol dependence in humans.

Various experimental methods are available for achieving the high blood alcohol levels (BALs) needed to induce alcohol dependence in rats, including gavage (Majchrowicz, 1975), diet (Lieber and DeCarli, 1982), intraperitoneal injections (Varlinskaya and Spear, 2004), and vapor exposure (Rogers et al., 1979). Vapor exposure allows the experimenter control over the amount, duration, and pattern of alcohol administration. Alcohol is administered passively through vapor delivered into the homecage air supply, decreasing the frequency of handling and/or restraint required for some other alcohol administration procedures. The experimenter controls the amount of alcohol administered to each cage, allowing for titration of BALs and achieving reasonably uniform intoxication levels among subjects in a given experiment. Rats exhibit physiological and behavioral signs of alcohol dependence after a few weeks. Behavioral testing can occur during acute withdrawal or protracted abstinence from alcohol vapor; the former allowing for repeated testing (e.g., within-subjects drug dose-response) at the same withdrawal time point on different days. In this protocol, we provide detailed, step-by-step methods updated from previous protocols (Gilpin et al., 2008) for induction of alcohol dependence using chronic intermittent exposure (CIE) to alcohol vapor in rats.

Materials and Reagents

  1. Razor blade (VWR, catalog number: 55411-055)
  2. 1.5 ml microtubes (Phenix Research Products, catalog number: MH-815)
  3. Rats (e.g., Wistar rats, 8 weeks of age; Charles River)
  4. 95% (v/v) ethanol (Decon Labs, Koptec, catalog number: V1105)
  5. Alcohol oxidase/buffer reagent (Analox Instruments, catalog number: GMRD-113)
  6. 100% (v/v) ethanol (Decon Labs, Koptec, catalog number: V1001G) 
  7. Liquinox (Alconox, catalog number: 1232)

Equipment

  1. Pre-assembled alcohol vapor inhalation system (La Jolla Alcohol Research, Inc.) or equivalent homemade system
  2. High-speed microcentrifuge (Fisher Scientific, AccuSpin Micro 17, catalog number: 13-100-675)
  3. Analox AM1 analyzer (Analox Instruments, P-GM7Micro-Stat, catalog number: Analox-AM1)
  4. Five-gallon carboy (Fisher Scientific, Thermo Scientific Nalgene Carboy, catalog number: 02-960-20A)
  5. Power strip with a timer (Century, model: BNC-U1)
  6. Animal scale (OHaus, Valor 3000 Xtreme, model: V31XH2)

Procedure

  1. Prepare vapor chamber by ensuring the pre-assembled air and vacuum lines and pressure gauges are securely connected to the air and vacuum lines for the vapor inhalation system (Figure 1).
  2. Connect ethanol delivery line to a five-gallon carboy containing 95% (v/v) ethanol and secure the cap (Figure 1A).
    Note: Do not use 100% ethanol, as it is toxic to animals, and do not use ethanol that is denatured or contains methyl ethyl ketone (MEK), denatonium benzoate, or benzonite.


    Figure 1. Assembled vapor inhalation system. A. Complete system. 95% (v/v) ethanol is delivered from a 5-gallon carboy reservoir (a) to the glass flask (b). A heating element is used to generate ethanol vapor, which is delivered to cages housed in the setup via incoming air lines. The temperature of the air delivered to the cages is not appreciably altered by this process. The amount of ethanol delivered through the air lines is controlled by the pump (c), which controls the frequency and volume of ethanol deliveries to the heated flask, where it is evaporated. Air flow is monitored via the air gauge (d). B. Close-up image of the pump. C. Close-up image of heating element, power source for heating element, and air gauge.

  3. Adjust pump settings to desired levels, based on animal weight and desired BALs.
    Note: These settings can be guided by a general rubric provided by the manufacturer. It is best to start at lower pump settings and gradually increase BALs over the course of the first week, rather than having rats start at a high BAL from session one (see Table 1 for an example). Further considerations for pump settings can be found in the Notes section of this protocol.
  4. Set the timer on the power strip to the desired settings. In our experiments, the ethanol pump (plugged into the power strip) is set to ON for 14 h a day, and OFF 10 h a day.
    Note: Vapor ON/OFF times can differ, but the experimental demands may dictate this schedule. The experimenter should be able to access animals at specific time points in the “vapor ON” and “vapor OFF” phases for measurement of BALs (at least once weekly, more often in the beginning of the exposure) and behavioral testing. For our experiments, we often set vapor ON at 6:00 PM and vapor OFF at 8:00 AM, with behavioral tests occurring at approximately 3:00 PM. Lights in our vapor room turn OFF at 8:00 AM, which means that most of the alcohol vapor exposure occurs during the light cycle and testing occurs during the dark cycle.
  5. Remove lid of the rats’ home cage, but keep the wire lid with food and water reservoirs in place. Insert cages into vapor inhalation system and secure the enclosure.
    Note: Never turn off incoming air to the vapor inhalation system when animals are inside the vapor inhalation system. If air pressure drops (detectable with the air gauge), remove cages immediately. When combining CIE with behavioral experiments that utilize repeatable procedures, it is ideal to collect baseline behavioral data (e.g., hindpaw withdrawal threshold using a Hargreaves apparatus for nociception, or lever pressing for operant self-administration studies; baseline data should be collected until individual behavior stabilizes to within 20% variability between sessions), and use those data to assign rats to “vapor” or “air” exposed groups that are counterbalanced for baseline behavioral data. Rats should be weighed before the first exposure to vapor and throughout the CIE procedure (e.g., twice per week).
  6. Perform vapor exposure, and monitor BALs. At the vapor OFF time point following the first session in vapor, monitor rats for intoxication by visually assessing somatic symptoms (e.g., gait, respiration, locomotion) while rats are in the homecage (e.g., Nixon and Crews, 2002). Using a razor blade, make a cut at the tip of the tail for blood collection; gently apply pressure to the length of the tail to encourage blood flow, and use a 1.5 ml microtube to collect blood from each animal (volume needed may differ based on the assay being used to measure BALs; for our experiments, we collect ~100 μl). This procedure can be performed with minimal restraint; place the rat on the experimenter’s non-dominant arm, encouraging the rat’s head to nestle in between the elbow and body, and use non-dominant hand to secure the rat’s tail. Place the rat’s tail on a sterile surface, and use the dominant hand to nick the tip of the tail. Return rats to homecage when finished collecting blood. Blood should also be collected from air-exposed controls to match for the stress of this procedure.
    Note: Microtubes containing heparin or other anticoagulant may be used; these should not interfere with BAL measurement using the Analox system (Step 7). Each time a change is made to alcohol pump settings, blood should be collected to measure BALs at that new setting. This procedure should be repeated until rats achieve the desired BAL range (in our work, typically 150-200 mg/dl). After the target BAL range is achieved, BALs should be measured at least once weekly thereafter to detect and account for metabolic tolerance and/or potential drift in settings. Adjustments will likely be needed as rats develop metabolic tolerance for alcohol following repeated exposures. If combining with behavioral experiments, perform tail bleeds on days when animals are not being tested. Air-exposed controls should be handled and bled on a schedule that is matched to alcohol vapor-exposed rats.
  7. Centrifuge blood samples at 9,500 x g for 12 min at room temperature to separate serum from blood. Determine BALs by running serum samples on an Analox AM1 analyzer, using a reconstituted alcohol oxidase/buffer reagent, according to the manufacturer’s instructions. An ethanol standard, used to calibrate the Analox prior to running serum samples, should be as close as possible to the anticipated range of BALs. In our experiments, we prepare a 100 mg/dl standard by adding 62.5 μl 100% (v/v) ethanol to 50 ml DI H2O. Standard solutions should be prepared frequently (at least every 1-2 weeks) to ensure accurate concentration.
    Note: If animals are extremely impaired (e.g., loss of locomotor activity, loss of muscle tone, no reaction to touch), this likely indicates extremely high BALs (can be confirmed by measuring BALs). If this exposure level is not the aim of the study, animals should be removed from alcohol vapor overnight to allow BALs to return to zero before the start of the next exposure. Consult with veterinary staff if additional intervention is necessary (e.g., saline administration if rat appears dehydrated; liquefied food if unable to eat solid food; placing on a heating pad if body temperature decreases; pharmacological treatment if severe withdrawal symptoms [e.g., seizures] are present). Adjust pump settings as necessary to reduce BALs. If unable to access an Analox AM1 analyzer, alternate methods for measuring BALs, such as commercially available enzymatic assays and gas chromatography-mass spectrometry, are available. If alternate methods are utilized, adjust the blood collection procedure (e.g., sample storage, centrifuge settings) as necessary. 
  8. Monitor rats throughout the CIE period. Assess rats daily for general health and well-being by visually observing behavior. Check food and water amounts daily, and refill when necessary. Replace bedding once weekly, on non-test days.
    Note: Vapor delivery to the cage generally does not result in wet food or bedding; in the event of condensation buildup, transfer animals to a clean, dry cage, and replace food and bedding. Withdrawal symptom severity can be assessed by the experimenter using desired metrics. While described in greater detail elsewhere (Majchrowicz, 1975; Faingold, 2008), symptoms that manifest during the withdrawal period, as BALs approach 0, can range from mild (e.g., general hyperactivity) to more severe (e.g., convulsive seizures). In the event of seizures, animals may be treated with a benzodiazepine or returned to alcohol vapor to relieve symptoms. It is best to consult with your institution’s veterinarian to develop an intervention plan prior to initiating experiments.
      A very rough estimation of the rate at which rats typically metabolize ethanol is approximately 25 mg/dl/h. Tests or tissue collection intended to access the intoxication or withdrawal stages should be scheduled accordingly (i.e., when BALs are peaking or equal to zero). For example, rats being tested for acute withdrawal effects might be maintained in a target range of 150-200 mg/dl and tested or sacrificed at 6-8 h after vapor OFF.
  9. Following completion of the experiment, turn the vapor inhalation system off. Clean cages with a mild detergent (e.g., Liquinox), and allow all components (cages and air/vacuum hoses) to completely dry.

Data analysis

Data analysis will typically differ based on the aims of the experiment and the in vivo and/or ex vivo procedures being used. If CIE is combined with behavioral testing, rats should be tested until a behavioral phenotype manifests (for example, escalated alcohol drinking and hyperalgesia emerge on the order of weeks; e.g., Somkuwar et al., 2016). The experimental aims will dictate the testing and end-point schedule, both in terms of the total duration of CIE, as well as in terms of the test or sacrifice timepoint relative to “vapor OFF.” In our experiments, rats are tested during the acute withdrawal period. Rats should always be exposed, treated and tested in parallel with air-exposed controls, which allows for statistical comparison of these two groups on all variables being tested (e.g., one-way or two-way ANOVA; for example, see Avegno et al., 2018). If CIE is used to prepare tissue for molecular biology or electrophysiology experiments, tissue harvesting often occurs after ≥ 4 weeks of CIE (with time = 0 corresponding to the first session in which rats exhibited BALs in the desired range). Correlational analyses can be used to explore the relationship between in vivo and/or ex vivo measures and BALs across animals in the same cohort.
  In behavioral experiments, rats may be excluded based on pre-determined criteria specific to the behavioral assay (e.g., failure to reach criteria, or following an outlier test). Furthermore, experimenters should be aware that extreme alcohol over-exposure (i.e., extremely high BALs) can produce neurotoxic effects (e.g., Obernier et al., 2002) and/or permanently alter behavior. The weight of each rat should be monitored throughout the experiment; weight loss of >10% may also be justification for removal of a subject from analysis (consult your veterinarian and IACUC), as this may indicate poor health outcome that can influence other variables of interest.

Notes

If desired, air-ethanol concentrations can be monitored with a breathalyzer apparatus. In order to achieve this, one only needs a port with a screw cap that can be removed to allow for air to escape into the breathalyzer. This port can be inserted by manufacturer, or by the experimenter in a self-assembled apparatus. Breathalyzers are available from multiple vendors. Typically, in our lab, we use blood ethanol concentrations (not air-ethanol concentrations) as the guide for setting changes. Sample settings for a machine with 4 rat cages and passive exhaust at the beginning of an experiment are shown in Table 1. Usually, these settings are too low to produce intoxication, which is intentional, and we slowly increase settings over days to produce intoxication. Settings would be different for a machine with more than 4 rat cages, or for animals of different size/age.
  Individual differences in BALs within a cohort can occur, due to differences in rat size and ethanol metabolism. In our experience with Long-Evans and Wistar rats, a given cohort will typically maintain BALs within a 50 mg/dl range of each other. Occasionally, considerable differences are observed, with a given rat exhibiting a BAL 100 mg/dL out of range of the others. Also, the concentration of alcohol vapor being delivered to each cage may differ slightly in some machines. Strategies to mitigate individual differences within a given cohort of rats will be dictated by the type of vapor machine being used. For example, if using a machine that delivers alcohol vapor to each cage using individual and separate lines, the experimenter may re-house animals (i.e., house a low BAL rat with a similarly low BAL cagemate) and/or change air flow to each individual cage. This would not be possible in a machine that delivers alcohol vapor to a single enclosed space where all animals are housed, but that type of machine should be subject to somewhat less inter-animal variability. Incremental increases or decreases in air flow (e.g., by ~3 p.s.i.) can produce slight decreases or increases in BALs, respectively. This strategy should be considered only when minor changes in individual BALs are desired, and air flow to any single standard size rat cage should never be below 10 p.s.i. More crude and sweeping changes in BALs can and should be achieved by changing alcohol pump settings.
  Rat strains may differ in terms of intoxication levels and metabolic tolerance in response to the same alcohol vapor settings (e.g., Gilpin et al., 2008). Age is also a consideration; for experiments intended to focus on adulthood, vapor exposure should begin no earlier than 8 weeks of age. Vapor exposure can occur at earlier timepoints (e.g., during adolescence), but interpretation of those data may be different from adult experimental data because alcohol effects on brain and behavior change across the lifespan (Novier et al., 2015). Experiments using adolescent rats may need to use lower pump settings to achieve target BALs, given their smaller size relative to adults; however, adolescent rats demonstrate an increased metabolism and decreased sensitivity to sedative effects of alcohol (Little et al., 1996). Sex may also be an important consideration, both in terms of body weight (female rats are smaller) and estrous cycle if that is an important consideration for the given experiment. CIE can also be used in different species, including the mouse, although settings and exposure schedule typically differ significantly (e.g., Becker and Hale, 1993; Griffin et al., 2009).

Table 1. Sample initial pump settings and corresponding BALs for a group of 8 male Wistar rats. Note that the amount of ethanol delivered to the heated flask at a given pump setting is different for each vapor system, and that pump settings on a given vapor system are not linear (i.e., incremental increases in pump settings do not necessarily translate to equally incremental increases in the amount of ethanol delivered to the system). This underscores the importance of measuring BALs each time an adjustment to the vapor system is made.

Acknowledgments

This protocol is supported by National Institute of Health (NIH) grants R01 AA023305 (NWG), R01 AA026531 (NWG), F32 AA025831 (EMA), and V.A. grant I01 BX003451 (NWG). The protocol outlined herein is adapted from Gilpin et al., 2008.

Competing interests

NWG owns shares in Glauser Life Sciences, Inc., a start-up company with interest in development of therapeutics for treatment of mental illness (no direct link to the current work). EMA declares no competing financial interests.

Ethics

All procedures were approved by the Institutional Animal Care and Use Committee of the Louisiana State University Health Sciences Center, and were in accordance with the National Institute of Health guidelines.

References

  1. Avegno, E. M., Lobell, T. D., Itoga, C. A., Baynes, B. B., Whitaker, A. M., Weera, M. M., Edwards, S., Middleton, J. W. and Gilpin, N. W. (2018). Central amygdala circuits mediate hyperalgesia in alcohol-dependent rats. J Neurosci 38(36): 7761-7773.
  2. Becker, H. C. and Hale, R. L. (1993). Repeated episodes of ethanol withdrawal potentiate the severity of subsequent withdrawal seizures: an animal model of alcohol withdrawal "kindling". Alcohol Clin Exp Res 17(1): 94-98.
  3. Egli, M., Koob, G. F. and Edwards, S. (2012). Alcohol dependence as a chronic pain disorder. Neurosci Biobehav Rev 36(10): 2179-2192.
  4. Faingold, C. L. (2008) The Majchrowicz binge alcohol protocol: an intubation technique to study alcohol dependence in rats. Curr Protoc Neurosci Chapter 9: Unit 9.28.
  5. Gilpin, N. W., Richardson, H. N., Cole, M. and Koob, G. F. (2008). Vapor inhalation of alcohol in rats. Curr Protoc Neurosci Chapter 9: Unit 9.29.
  6. Griffin, W. C., 3rd, Lopez, M. F., Yanke, A. B., Middaugh, L. D. and Becker, H. C. (2009). Repeated cycles of chronic intermittent ethanol exposure in mice increases voluntary ethanol drinking and ethanol concentrations in the nucleus accumbens. Psychopharmacology (Berl) 201(4): 569-580.
  7. Kallupi, M., Vendruscolo, L. F., Carmichael, C. Y., George, O., Koob, G. F. and Gilpin, N. W. (2014). Neuropeptide YY(2)R blockade in the central amygdala reduces anxiety-like behavior but not alcohol drinking in alcohol-dependent rats. Addict Biol 19(5): 755-757.
  8. Koob, G. F. (2003). Alcoholism: allostasis and beyond. Alcohol Clin Exp Res 27(2): 232-243.
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  10. Little, P. J., Kuhn, C. M., Wilson, W. A. and Swartzwelder, H. S. (1996). Differential effects of ethanol in adolescent and adult rats. Alcohol Clin Exp Res 20(8): 1346-1351.
  11. Majchrowicz, E. (1975). Induction of physical dependence upon ethanol and the associated behavioral changes in rats. Psychopharmacologia 43(3):245-254.
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  13. Novier, A., Diaz-Granados, J. L. and Matthews, D. B. (2015). Alcohol use across the lifespan: An analysis of adolescent and aged rodents and humans. Pharmacol Biochem Behav 133: 65-82.
  14. Obernier, J. A., Bouldin, T. W. and Crews, F. T. (2002). Binge ethanol exposure in adult rats causes necrotic cell death. Alcohol Clin Exp Res 26(4): 547-557.
  15. Roberts, A. J., Cole, M. and Koob, G. F. (1996). Intra-amygdala muscimol decreases operant ethanol self-administration in dependent rats. Alcohol Clin Exp Res 20(7): 1289-1298.
  16. Rogers, J., Wiener, S. G. and Bloom, F. E. (1979). Long-term ethanol administration methods for rats: advantages of inhalation over intubation or liquid diets. Behav Neural Biol 27(4): 466-486.
  17. Sacks, J. J., Gonzales, K. R., Bouchery, E. E., Tomedi, L. E. and Brewer, R. D. (2015). 2010 national and state costs of excessive alcohol consumption. Am J Prev Med 49(5): e73-e79.
  18. SAMHSA. (2015). Substance Abuse and Mental Health Services Administration (SAMHSA). 2015 National Survey on Drug Use and Health (NSDUH). Table 5.6A–Substance Use Disorder in Past Year among Persons Aged 18 or Older, by Demographic Characteristics: Numbers in Thousands, 2014 and 2015. Available at https://www.samhsa.gov/data/sites/default/files/NSDUH-DetTabs-2015/NSDUH-DetTabs-2015/NSDUH-DetTabs-2015.htm#tab5-6a.
  19. Somkuwar, S. S., Fannon, M. J., Staples, M. C., Zamora-Martinez, E. R., Navarro, A. I., Kim, A., Quigley, J. A., Edwards, S. and Mandyam, C. D. (2016). Alcohol dependence-induced regulation of the proliferation and survival of adult brain progenitors is associated with altered BDNF-TrkB signaling. Brain Struct Funct 221(9): 4319-4335. 
  20. Varlinskaya, E. I. and Spear, L. P. (2004). Acute ethanol withdrawal (hangover) and social behavior in adolescent and adult male and female Sprague-Dawley rats. Alcohol Clin Exp Res 28(1): 40-50.
  21. Walker, B. M. and Koob, G. F. (2007). The gamma-aminobutyric acid-B receptor agonist baclofen attenuates responding for ethanol in ethanol-dependent rats. Alcohol Clin Exp Res 31(1): 11-18.

简介

【摘要】酒精使用障碍(AUD)是一种重要的公共健康和经济负担,其特征通常是反复发作酒精中毒和戒断。 事实上,所有器官系统都受到慢性酒精暴露的影响。 可以使用大鼠作为模型生物来研究这些效果; 然而,大鼠通常不会将酒精自我施用到达到依赖的生理和行为方面所必需的水平。 本文所述的方案可用于通过向肠笼中长时间施用酒精蒸气来诱导大鼠的酒精依赖。 该方法允许研究人员控制中毒的水平,持续时间和模式,并且可靠地诱导酒精依赖的生理和行为方面,允许研究与人类中的AUD相关的生物学和行为。

【背景】酒精使用障碍(AUD)影响超过1500万美国人(SAMHSA,2015),每年花费美国约2490亿美元(Sacks et al。>,2015)。澳元的逐步发展的特点是反复进行酒精中毒和戒断,这引发了一系列神经适应和行为改变。有AUD的人饮用过量的酒精,部分原因是酒精含有阳性(即>,饮酒以获得奖励效果)和消极(即>,饮酒以消除或避免厌恶的感觉)加强对这些人的影响(Koob,2003)。戒酒的酒精依赖大鼠表现出酒精依赖人类表现出的许多相同行为,包括饮酒量的增加(例如>,Roberts 等。>,1996),获得酒精的动机(例如>,Walker和Koob,2007),类似焦虑的行为(例如>,Kallupi et al。>,2014)和伤害感受( eg >,Egli et al。>,2012; Avegno et al。>。,2018)。这些行为表明,大鼠酒精依赖模型是研究人类酒精依赖的神经生物学基础的有用工具。

各种实验方法可用于实现诱导大鼠酒精依赖所需的高血液酒精水平(BAL),包括灌胃(Majchrowicz,1975),饮食(Lieber和DeCarli,1982),腹膜内注射(Varlinskaya和Spear,2004),和蒸气暴露(Rogers et al。>。,1979)。蒸汽暴露允许实验者控制酒精施用的量,持续时间和模式。通过输送到母笼空气供应的蒸汽被动地施用酒精,降低了一些其他酒精施用程序所需的处理和/或限制的频率。实验者控制给予每个笼子的酒精量,允许滴定BAL并在给定实验中实现受试者中合理均匀的中毒水平。几周后,大鼠表现出酒精依赖的生理和行为迹象。行为测试可在急性戒断或长期戒酒期间发生;前者允许在不同日期的相同戒断时间点重复检测(例如>,受试者内药物剂量 - 反应)。在该协议中,我们提供了从先前的方案(Gilpin 等人>,2008)更新的详细的逐步方法,用于使用慢性间歇暴露(CIE)对大鼠酒精蒸气诱导酒精依赖。

关键字:酒精依赖, 酒精使用障碍, 蒸汽, 脱瘾, 中毒, 血液酒精水平, 大鼠

材料和试剂

  1. 剃刀刀片(VWR,目录号:55411-055)
  2. 1.5 ml microtubes(Phenix Research Products,目录号:MH-815)
  3. 大鼠(例如>,Wistar大鼠,8周龄;查尔斯河)
  4. 95%(v / v)乙醇(Decon Labs,Koptec,目录号:V1105)
  5. 醇氧化酶/缓冲试剂(Analox Instruments,目录号:GMRD-113)
  6. 100%(v / v)乙醇(Decon Labs,Koptec,目录号:V1001G) 
  7. Liquinox(Alconox,目录号:1232)

设备

  1. 预装酒精蒸气吸入系统(La Jolla Alcohol Research,Inc。)或等效的自制系统
  2. 高速微量离心机(Fisher Scientific,AccuSpin Micro 17,目录号:13-100-675)
  3. Analox AM1分析仪(Analox Instruments,P-GM7Micro-Stat,目录号:Analox-AM1)
  4. 五加仑carboy(Fisher Scientific,Thermo Scientific Nalgene Carboy,目录号:02-960-20A)
  5. 带定时器的电源板(世纪,型号:BNC-U1)
  6. 动物秤(OHaus,Valor 3000 Xtreme,型号:V31XH2)

程序

  1. 通过确保预组装的空气和真空管路和压力表牢固地连接到蒸汽吸入系统的空气和真空管路来准备蒸汽室(图1)。
  2. 将乙醇输送管线连接到含有95%(v / v)乙醇的5加仑汽车,并固定盖子(图1A)。
    注意:不要使用100%乙醇,因为它对动物有毒,并且不使用变性的乙醇或含有甲基乙基酮(MEK),苯甲酸地那铵或苯甲酸盐。>


    图1.组装蒸汽吸入系统。 A.完整系统。将95%(v / v)乙醇从5加仑的carboy储器(a)输送到玻璃烧瓶(b)。加热元件用于产生乙醇蒸汽,其通过进入的空气管线输送到容纳在装置中的笼子。输送到笼子的空气温度不会因此过程而明显改变。通过空气管线输送的乙醇量由泵(c)控制,泵(c)控制乙醇输送到加热烧瓶的频率和体积,在那里蒸发。通过空气压力计(d)监测空气流量。 B.泵的特写图像。 C.加热元件的特写图像,加热元件的电源和气压计。

  3. 根据动物体重和所需BAL将泵设置调整到所需水平。
    注意:这些设置可以由制造商提供的一般规则指导。最好从较低的泵设置开始,并在第一周的过程中逐渐增加BAL,而不是让大鼠从第一阶段的高BAL开始(参见表1的例子)。有关泵设置的更多注意事项,请参阅本协议的“注释”部分。>
  4. 将电源板上的计时器设置为所需的设置。在我们的实验中,乙醇泵(插入电源板)每天开启14小时,每天关闭10小时。
    注意:蒸汽开/关时间可能不同,但实验要求可能决定了这个时间表。实验者应该能够在“蒸汽开”和“蒸汽关闭”阶段的特定时间点接近动物以测量BAL(至少每周一次,更经常在暴露开始时)和行为测试。对于我们的实验,我们经常在下午6:00将蒸汽设置为开启,并在上午8:00设置蒸汽关闭,行为测试在下午3:00左右进行。我们的蒸汽室中的灯在上午8:00关闭,这意味着大部分酒精蒸气暴露发生在灯光循环期间,测试发生在黑暗循环期间。>
  5. 取下老鼠家笼的盖子,但保持电线盖上有食物和水库。将保持架插入蒸汽吸入系统并固定外壳。
    注意:当动物在蒸汽吸入系统内时,从不关闭进入蒸汽吸入系统的空气。如果气压下降(可通过气压计检测到),请立即取下保持架。当将CIE与利用可重复程序的行为实验相结合时,理想的是收集基线行为数据(> 例如> ,使用Hargreaves装置进行伤害感受的后爪退缩阈值,或杠杆按压操作性自我管理研究;应收集基线数据,直到个体行为稳定到会话之间的变异性在20%之内),并使用这些数据将大鼠分配给基线行为数据平衡的“蒸气”或“空气”暴露组。在第一次暴露于蒸气之前和整个CIE程序(> 例如> ,每周两次)之前,应对大鼠进行称重。>
  6. 进行蒸汽暴露,并监测BAL。在蒸汽第一次训练后的蒸汽关闭时间点,通过目视评估大鼠在家中的躯体症状(例如>,步态,呼吸,运动)来监测大鼠中毒(例如< / em>,Nixon and Crews,2002)。使用剃刀刀片,在尾巴尖端切开血液进行采血;轻轻地向尾巴长度施加压力以促进血液流动,并使用1.5 ml微管从每只动物采集血液(根据用于测量BAL的测定,所需的体积可能不同;对于我们的实验,我们收集~100μl )。这个程序可以在最小的限制下进行;将大鼠放在实验者的非优势手臂上,鼓励大鼠的头部位于肘部和身体之间,并使用非惯用手固定老鼠的尾巴。将大鼠的尾巴放在无菌的表面上,并用惯用的手来划伤尾巴的尖端。收集血液后,将老鼠送回家中。还应从暴露在空气中的对照中采集血液,以配合此程序的压力。
    注意:可以使用含有肝素或其他抗凝血剂的微管;这些不应该干扰使用Analox系统的BAL测量(步骤7)。每次更改酒精泵设置时,都应收集血液以测量新设置的BAL。应重复该过程,直到大鼠达到所需的BAL范围(在我们的工作中,通常为150-200mg / dl)。在达到目标BAL范围后,应该每周至少测量一次BAL以检测并解释设置中的代谢耐受性和/或潜在漂移。由于大鼠在反复接触后会对酒精产生代谢耐受,因此可能需要进行调整。如果与行为实验相结合,在动物未进行测试的日子进行尾部出血。空气暴露的对照应按照与暴露于酒精蒸气的大鼠相匹配的时间表进行处理和放血。>
  7. 在室温下以9,500 x g >离心血液样品12分钟以从血液中分离血清。根据制造商的说明,使用重组的醇氧化酶/缓冲试剂,在Analox AM1分析仪上运行血清样品,以确定BAL。用于在运行血清样品之前校准Analox的乙醇标准品应尽可能接近预期的BAL范围。在我们的实验中,我们通过向50ml DI H 2 中加入62.5μl100%(v / v)乙醇制备100mg / dl标准品。应经常(至少每1-2周)准备标准溶液,以确保准确浓度。
    注意:如果动物极度受损(> 例如> ,失去运动活动,失去肌肉张力,没有反应),这可能表明BALs极高(可以通过测量BAL来确认。如果该暴露水平不是研究的目的,应将动物从酒精蒸汽中过夜除去,以使BAL在下一次暴露开始前恢复到零。如果需要进一步干预,请咨询兽医人员(> 例如> ,如果大鼠出现脱水则给予盐水;如果不能吃固体食物则给液化食物;如果体温则放在加热垫上减少;如果存在严重的戒断症状[> 例如> ,癫痫发作],则进行药物治疗。根据需要调整泵设置以减少BAL。如果无法使用Analox AM1分析仪,可以使用其他方法测量BAL,例如市售的酶测定和气相色谱 - 质谱。如果使用其他方法,请根据需要调整血液采集程序(> 例如> ,样品存储,离心机设置)。>&nbsp;
  8. 在整个CIE期间监测大鼠。通过视觉观察行为,每天评估大鼠的一般健康状况和健康状况。每天检查食物和水量,必要时补充。在非测试日,每周更换一次床上用品。
    注意:向笼子输送蒸汽通常不会导致潮湿的食物或床上用品;如果出现结露,将动物转移到干净,干燥的笼子中,并更换食物和床上用品。实验者可以使用期望的度量来评估戒断症状严重性。虽然在其他地方有更详细的描述(Majchrowicz,1975; Faingold,2008),但在退出期间出现的症状,如BAL接近0,可以是温和的(> 例如> ,一般多动症)更严重(> 例如> ,惊厥性癫痫发作)。在癫痫发作的情况下,动物可以用苯二氮卓类药物治疗或返回酒精蒸气以缓解症状。在开始实验之前,最好咨询您所在机构的兽医,制定干预计划。 >
    &NBSP;对大鼠通常代谢乙醇的速率的非常粗略的估计是大约25mg / dl / h。用于进入中毒或戒断阶段的测试或组织收集应相应地安排(> 即> ,当BAL达到峰值或等于零时)。例如,正在测试急性戒断效应的大鼠可能会维持在150-200 mg / dl的目标范围内,并在蒸汽关闭后6-8小时进行测试或处死。>
  9. 完成实验后,关闭蒸汽吸入系统。使用温和的清洁剂(例如>,Liquinox)清洁笼子,并让所有组件(笼子和空气/真空软管)完全干燥。

数据分析

数据分析通常基于实验的目的和使用的体内>和/或离体>程序而不同。如果将CIE与行为测试相结合,则应对大鼠进行测试,直至出现行为表型(例如,升级的酒精饮酒和痛觉过敏出现在几周; 例如>,Somkuwar 等。< / em>,2016)。实验目标将根据CIE的总持续时间以及相对于“蒸汽关闭”的测试或牺牲时间点来规定测试和终点时间表。在我们的实验中,大鼠在测试期间进行测试。急性停药期。大鼠应始终暴露,处理和与暴露于空气的对照平行测试,这允许对所测试的所有变量进行这两组的统计比较(例如>,单向或双向ANOVA;例如,参见Avegno et al。>,2018)。如果CIE用于制备用于分子生物学或电生理学实验的组织,则组织收获通常在≥4周的CIE后发生(时间= 0对应于第一阶段,其中大鼠表现出所需范围内的BAL)。相关分析可用于探索同一队列中动物体内体内>和/或体外>测量值与BALs之间的关系。
&NBSP;在行为实验中,可以基于行为测定特异性的预定标准(例如>,未达到标准,或在异常值测试之后)排除大鼠。此外,实验者应该意识到极端酒精过度暴露(即>,极高的BALs)会产生神经毒性作用(例如>,Obernier 等。 >,2002)和/或永久改变行为。在整个实验过程中应监测每只大鼠的体重;体重减轻> 10%也可能是从分析中去除受试者的理由(咨询您的兽医和IACUC),因为这可能表明可能影响其他感兴趣变量的健康结果不佳。

笔记

如果需要,可以用呼气测醉装置监测空气 - 乙醇浓度。为了达到这个目的,人们只需要一个带螺帽的端口,可以将其移除以允许空气进入呼吸测醉器。该端口可由制造商插入,或由实验者插入自组装设备中。多个供应商提供呼吸测醉器。通常,在我们的实验室中,我们使用血液乙醇浓度(不是空气 - 乙醇浓度)作为设定变化的指南。表1中显示了在实验开始时具有4个鼠笼和被动排气的机器的样品设置。通常,这些设置太低而不能产生中毒,这是有意的,并且我们在数天内缓慢增加设置以产生中毒。对于具有4个以上鼠笼的机器或不同大小/年龄的动物,设置会有所不同。
&NBSP;由于大鼠大小和乙醇代谢的差异,群体内BAL的个体差异可能发生。根据我们对Long-Evans和Wistar大鼠的经验,给定的队列通常将BAL保持在彼此50mg / dl范围内。偶尔会观察到相当大的差异,给定的大鼠显示BAL 100 mg / dL超出其他大鼠的范围。此外,在一些机器中,输送到每个笼子的酒精蒸汽浓度可能略有不同。减轻给定队列大鼠内个体差异的策略将取决于所用蒸汽机的类型。例如,如果使用通过单独和单独的线路向每个笼子输送酒精蒸汽的机器,实验者可以重新安置动物(即>,容纳具有类似低BAL笼状物的低BAL大鼠)和/或改变每个笼子的气流。这在将酒精蒸汽输送到所有动物所在的单个封闭空间的机器中是不可能的,但是这种类型的机器应该在较小的动物间变异性。空气流量的增量增加或减少(例如>,~3 p.s.i.)可分别使BAL略微减少或增加。只有在需要对个体BAL进行微小改变时才应考虑该策略,并且任何单个标准尺寸鼠笼的气流量不应低于10 p.s.i.通过改变酒精泵的设置,可以而且应该通过更加粗暴和彻底的BAL变化来实现。
&NBSP;响应于相同的醇蒸气设置,大鼠菌株在中毒水平和代谢耐受性方面可能不同(例如>,Gilpin 等人>,2008)。年龄也是一个考虑因素;对于旨在关注成年期的实验,蒸气暴露应该不早于8周龄开始。蒸汽暴露可能发生在较早的时间点(例如>,在青春期),但这些数据的解释可能与成人实验数据不同,因为酒精对大脑和行为的影响在整个生命周期中发生变化(Novier et al。>,2015)。使用青少年大鼠的实验可能需要使用较低的泵设置来实现目标BAL,因为它们相对于成人的尺寸较小;然而,青春期大鼠表现出新陈代谢增加和对酒精镇静作用的敏感性降低(Little et al。>,1996)。性别也可能是一个重要的考虑因素,无论是体重(雌性大鼠体重较小)还是动情周期,如果这是给定实验的重要考虑因素。 CIE也可以用于不同的物种,包括小鼠,虽然设置和暴露时间表通常差别很大(例如>,Becker和Hale,1993; Griffin et al。>,2009 )。

表1.一组8只雄性Wistar大鼠的初始泵设置和相应BAL的样本。请注意,在给定泵设置下输送到加热烧瓶的乙醇量对于每个蒸汽系统是不同的,并且给定蒸汽系统上的泵设置不是线性的(即>,泵设置的增量增加不一定转化为输送到系统的乙醇量的同等增量增加)。这强调了每次调整蒸汽系统时测量BAL的重要性。

致谢

该方案得到国立卫生研究院(NIH)授予的R01 AA023305(NWG),R01 AA026531(NWG),F32 AA025831(EMA)和V.A.的支持。授予I01 BX003451(NWG)。本文概述的方案改编自(Gilpin 等人,>,2008)。

利益争夺

NWG拥有Glauser Life Sciences,Inc。的股份,该公司是一家有兴趣开发精神疾病治疗药物的初创公司(与当前工作没有直接联系)。 EMA声明没有竞争性的经济利益。

伦理

所有程序均经路易斯安那州立大学健康科学中心的机构动物护理和使用委员会批准,并符合国家卫生研究院的指导方针。

参考

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Copyright: © 2019 The Authors; exclusive licensee Bio-protocol LLC.
引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. Avegno, E. M. and Gilpin, N. W. (2019). Inducing Alcohol Dependence in Rats Using Chronic Intermittent Exposure to Alcohol Vapor. Bio-protocol 9(9): e3222. DOI: 10.21769/BioProtoc.3222.
  2. Avegno, E. M., Lobell, T. D., Itoga, C. A., Baynes, B. B., Whitaker, A. M., Weera, M. M., Edwards, S., Middleton, J. W. and Gilpin, N. W. (2018). Central amygdala circuits mediate hyperalgesia in alcohol-dependent rats. J Neurosci 38(36): 7761-7773.
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