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Dec 2017

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Behavioral Evaluation of Seeking and Preference of Alcohol in Mice Subjected to Stress
承受应激小鼠的觅食和酒精偏好的行为学评估   

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Abstract

The alcohol preference model is one of the most widely used animal models relevant to alcoholism. Stressors increase alcohol consumption. Here we present a protocol for a rapid and useful tool to test alcohol preference and stress-induced alcohol consumption in mice. In this model, animals are given two bottles, one with a diluted solution of ethanol in water, and the other with tap water. Consumption from each bottle is monitored over a 24-h period over several days to assess the animal’s relative preference for the ethanol solution over water. In the second phase, animals are stressed by restraining them for an hour daily and their subsequent preference of tap water or the ethanol solution is evaluated. Preference is measured by the volume and/or weight or liquid consumed daily, which is then converted to a preference ratio. The alcohol preference model was combined with the conditioned place preference paradigm to determine alcohol conditioning and preference following the deletion of CB2 cannabinoid receptors in dopaminergic neurons in the DAT-Cnr2 Cre-recombinant conditional knockout (cKO) mice in comparison with the wild-type control mice.

Keywords: Alcohol (酒精), Stress (应激), Mouse model (小鼠模型), Behavior (行为), Cannabinoid (大麻素), Conditioned place preference (大麻素位置偏爱)

Background

Many aspects of alcoholism and alcohol consumption can be studied through animal models. Alcohol induces positive reinforcement, and animals can seek alcohol and even work for it. However, alcohol can also be a negative reinforcement, since it is capable of reducing anxiety. No animal model is able to duplicate the complex features of alcoholism. Oral ethanol self-administration is widely used for examining specific aspects of behavior and physiology relevant for understanding alcoholism (Mardones and Segovia-Riquelme, 1983; Cunningham et al., 2000). Mice can be genetically manipulated at cell type specific levels and therefore are valuable for research into the cell type specific genetic determinants of alcoholism.

The alcohol preference model is one of the most widely used animal models relevant to alcoholism. This model meets important criterion, which is that the ethanol should be self-administered orally (Cicero, 1980; Crabbe et al., 2010). An animal’s genotype exerts a strong influence on self-administration in this model. Some mouse strains, like the inbred strain of mouse C57BL/6J (Rhodes et al., 2005), present a genetically influenced high preference for ethanol and they voluntarily consume it orally (Yoneyama et al., 2008; Barkley-Levenson and Crabbe, 2012). The conditioned place paradigm (CPP) is widely used to explore the effects of addictive substances including alcohol, taking advantage of learned associations. Therefore, alcohol CPP measures the association of alcohol with a particular environment to determine whether mice can acquire alcohol CPP.

Stress can interact with ongoing ethanol consumption to trigger increased intake (e.g., self-medicating behavior), thereby increasing initial susceptibility to alcohol use disorders. Among the stressors, a restraint model of acute and chronic stress can increase ethanol consumption (Yang et al., 2008).

New advances and accumulating evidence support a role for the endocannabinoid system in the effects of alcohol. The endocannabinoid system consists of two cannabinoid receptors, CB1Rs and CB2Rs, with endocannabinoids and the enzymes for the biosynthesis and inactivation of the endocannabinoids. Our goal here was to summarize the protocol used to measure alcohol preference in combination with stress-induced alcohol consumption. We also provide evidence that the endocannabinoid system plays a role in alcohol preference following dopaminergic neuron specific deletion of CB2Rs in the mouse model (Liu et al., 2017).

Materials and Reagents

  1. 50 ml Polypropylene Centrifuge Tubes with Attached Caps (Boekel Scientific, catalog number: 120021 )
  2. Bottles (see Figures 1B and 1C) with Sipper Caps (Chewy, catalog number: 101445 )
  3. Mice: Adult (7 weeks or older) mice (C57BL/6J) (THE JACKSON LABORATORY, catalog number: 000664 )
    Note: Alternate strains and ages of mice may also be used. Mice are housed alone, each in their respective cages, in an environment with controlled temperature (around 23 °C) and humidity under a 12-12 h light-dark cycle with free access to food. See Animal considerations in Notes for more details.
  4. 100% Alcohol and dilutions: 8%, 16% and 32% (Sigma-Aldrich, catalog number: 1012768 )


    Figure 1. Photos showing experimental set-up for the evaluation of alcohol preference. A. Tubes in which naïve or conditional knockout mice and wild-type controls are subjected to acute stress. B. Tubes, water bottles, and a clean empty mouse cage. C. Image of the control cage. D. Image of the apparatus for conditioned place preference.

Equipment

  1. Mouse polycarbonate home cages (7.5 in W x 11 in L x 5 in H) with standard woodchip mouse bedding (Fisher Scientific, catalog number: 01-286-13A)
    Manufacturer: Tecniplast, catalog number: 1290D00SU .
    Note: Standard cage changes are allowed during the habituation period (see below). However, it is recommended to avoid cage changes during the period of data collection to prevent leakage. The cage can be cleaned between phases. One cage for each subject mouse.
  2. Stainless steel wire cage lid modified to allow space for two bottles and the food (Fisher Scientific, catalog number: 01-286-13A)
    Manufacturer: Tecniplast, catalog number: 1290D00SU .
  3. Drill with small (1 mm) drill bit (for making three small holes in top of 50 ml Polypropylene Centrifuge Tubes, to allow the mouse to breathe, and a single larger hole in the cap to insert the tail)
  4. Thermo Scientific weighing scale (Thermo Fisher Scientific, catalog number: 20031 )

Software

  1. Activity Monitor software (Med Associates, St. Albans, VT) (for automated data collection)
  2. GraphPad PRISM 6.0 software (GraphPad Software. Inc., San Diego, CA, USA) (for data analysis)

Procedure

All experiments were conducted during dark phase, under dim light illumination.

  1. Experimental setup
    1. Animals: Naïve C57Bl/6Js were used, and as wild-type controls for the DAT-Cnr2 conditional knockout (cKO) mice. The generation and breeding of the cKO mice was previously reported (Liu et al., 2017).
    2. Experimental space should be quiet and dimly lit, under controlled humidity and temperature.
    3. All the cages should be placed on a rack designated for this experiment. The rack should be stable to avoid movement and to prevent leakage. The experimenter should be careful when removing the bottles to minimize the amount of leakage. Leakage should be avoided to reduce undesired variability.
    4. The experimental area should be clean, and all the measurements should be conducted at the same time of the day.
    5. One of the cages will be the “control cage”. This cage will have everything described above except a mouse.
    6. All cages should be labeled with the information for the specific experiment, such as the mouse’s strain, age, and gender. Food should always be kept on the right side for consistency.

  2. Setting up 2-bottle choice test
    1. Fill all the bottles with 150 ml of water and put on the sipper tap.
      Note: Make sure there are no water leaks by inverting the bottles.
    2. Label each bottle with the number of the subject mouse (1, 2, 3…) in permanent marker. Label the cages with the same number.
    3. Place the 2 bottles onto the wire lid (Figure 2), with minimal shaking to avoid dripping. Both bottles should be placed on one side of the divided wire rack, leaving the other side for food. Put food pellets on the other side of the wire lid.
      Note: Make sure that the food is always on the right side, to be consistent. Mice were adapted to the testing room, with the same conditions as in the colony room (22 ± 2 °C; humidity: 55 ± 5%; 12 h dim/artificial light/12 h dark cycle: light off at 7 PM).

  3. Habituation to the 2-bottle
    1. Place a single mouse in the proper mouse caging (see Equipment) under controlled conditions (see Materials and Reagents).
    2. Isolate the subjects into individual cages and give them two bottles filled with 150 ml water. Allow mice to acclimate for at least 4 days under these conditions.

  4. Setting up 2 bottle choice for alcohol preference
    1. Remove the bottles and clean them.
    2. Mix water with 100% alcohol to 16%.
    3. For each mouse, fill one bottle with 150 ml of water and another bottle with alcohol solution. Put the sipper cap on the bottle.
      Note: Make sure there are no water leaks by inverting the bottles.
    4. Label each bottle with the assigned number of the mouse, and water or alcohol in permanent marker (e.g., 1 H2O/1 EtOH).
    5. Record weight with cap on the bottle and write down the value on a record sheet.

  5. Evaluation of alcohol preference
    1. Place the 2 bottles onto the wire lid, with minimal shaking to avoid dripping. Both bottles should be placed on one side of the divided wire rack, leaving the other side for food. Put food pellets on the other side of the wire lid.
      Note: Make sure that the food is always on the right side, to be consistent.
    2. Assess water and alcohol solution consumption daily for 5 days. Record weight. Monitor for any problems with liquid flow or spillage, e.g., wet bedding below the sipper tube. Switch positions (left vs. right) of bottles daily.
    3. The “control cage” has just two bottles of water without a mouse. These bottles are weighed to determine the amount of spillage when bottles are inverted, and to later correct the data of the other groups.

  6. Acute stress-induced alcohol preference (5 days)
    1. Set up 2 bottle choices for alcohol preference (described above).
    2. After removing the bottles for daily weighing on the first day, mice are randomly assigned to be in the experimental or control group. Label the cages with “control” or “experimental” (above where the mouse’s number is) and place them by group on the shelf. The experimental group of mice are restrained in a 50 ml conical tube for an hour. After that, they are released and the bottles for both groups (control and experimental) and placed in the cages again.
    3. Repeat these steps for 5 consecutive days. The bottles are weighed each day, and the mice are weighed before and after each phase.
    4. The general timeline of the alcohol preference is depicted from Procedures D-F over the 5-day period. 

  7. Chronic mild stress (CMS) induced alcohol consumption (from 4-7 weeks) (Table 1)
    1. Set up separate groups of the wild-type and selected cKO mice (n = 6 per group) for experimental and control groups in separate rooms. All animals are housed individually, except for the non-stressed control groups.
    2. For the CMS regime, mice are subjected to various stressors according to a semi-random schedule Table 1 for 4-7 weeks as planned.
    3. The stress regime for each week consisted of food deprivation for 12 h, water deprivation for 12 h, damp bedding for 12 h, overnight stroboscopic illumination, tail suspension for 10 min, tube restrain for 30 min, as described above, loud music overnight, wire mesh to replace bedding 12 h, 30 min introduction of an intruder mouse to the cage and lights off or on. These stressors were paired morning or overnight.
    4. All non-CMS groups in the separate room were given food and water at all times.

  8. Conditioned place preference (CPP)
    Alcohol CPP training and testing are conducted using an infra-red photobeam detector open field apparatus (ENV-510) from Med Associates (St. Albans, VT, USA).
    1. Alcohol CPP training and testing are conducted using an infra-red photobeam detector open field apparatus (ENV-510) from Med Associates (St. Albans, VT, USA) equipped with the two-compartment place preference inserts (ENV-512), as shown in Figure 1D. The floor of compartment-1 has parallel rods (3-mm radius, 8 mm center to center spacing) with black cardboard paper covering the outside. Compartment-2 has a stainless steel wire mesh (6 x 6) floor with white cardboard paper covering the outside of the walls. Each compartment is 13 cm (width) x 15 cm (depth).
    2. Perform all conditioning sessions and preference tests in each group in three phases (Figure 3) (pre-conditioning, conditioning and post-conditioning phases).
    3. During the post-conditioning phase on the test day, mice are allowed to explore both compartments for 15 min.
    4. The CPP score is defined as the time spent in the alcohol paired compartment minus the time spent in the saline-paired compartment during the CPP test.


      Figure 2. Setup of 2 bottle choice for alcohol preference and water is available


      Figure 3. Graphical presentation of the CPP paradigm used. All conditioning sessions and preference tests were performed at the same time daily. Animals were allowed to habituate in the CPP apparatus for 45 min before the conditioning phase.

    Table 1. Chronic Mild Stressors (from 4-7 weeks) 

Data analysis

All data are transcribed in a spreadsheet. Set up a column for the raw weight data and next to it a column for the corrected weight data. The corrected weight data is the amount of spillage (of water and alcohol) each day from the control cage. This number will be subtracted from the raw weight data. Each day, the spillage amount must be subtracted from the raw water and alcohol weight data to determine the corrected weight.
  To calculate the amount of alcohol or water consumed per day, the bottle weights were averaged for each day. These values were then subtracted from the averages of the bottle weights for each group on that day. Subtracting the new bottle weight from that of the previous day gave the amount of alcohol or water drank by the mice overnight.
  Day 0 is when the bottles were initially filled with 150 ml of water/alcohol and therefore the consumption for that day was 0.
  The alcohol preference ratio was determined by dividing the amount of alcohol consumed by total liquid (alcohol + water) consumed.
  The activity monitor recorded time spent in each of the compartments, using the photo electric beam break counts. The time that each mouse spent in the compartment was recorded and the CPP score was defined as the time spent in the alcohol paired compartment minus the time spent in the saline-paired compartment during the CPP test. The activity monitor software (Med Associates, St. Albans, VT) was used for automated data collection.

Representative data: Figure 4 is representative data for Alcohol Preference Test. In this test, the mice were individually housed and habituated with two fluid bottles available for three days. We used two groups: 12 male C57/6J mice as wild-type and 12 male DAT-Cnr2 conditional knockout (cKO) mice that do not express the CB2R in midbrain dopamine neurons. On the day of the experiment, all the animals were weighed. One bottle was filled with 150 ml of tap water and the other was filled with 150 ml of 16% alcohol solution. They were weighed with the tops on and placed on top of the cages (labeled correctly). For the alcohol preference test, the bottles were weighed for each animal for five consecutive days at 10 AM. The positions of the bottles in the different cages were randomized with respect to which side of the cages they were placed. In the second part of the experiment, each group was then split into two groups. One group (n = 6) was stressed by putting them in a 50 ml tube for one hour each day for 5 consecutive days. The rest of the animals remained in their cages. All the bottles were weighed for each animal for five consecutive days at the same time, and all the bottles were placed on top of the cages at 11 AM. In all the experiments, the ratio of alcohol to water consumed and the total fluid consumption were calculated to obtain a preference ratio. The alcohol preference ratio was determined by dividing the amount of alcohol consumed by total fluid (alcohol + water) consumed, with and without the sub-acute stress. The wild-type mice preferred alcohol over water, as evident by the significant increase (as determined by Repeated Measurement ANOVA). The DAT-Cnr2 cKO mice (mice did not show an alcohol-CPP induction, Figure 5), and do not show a significant increase of the preference for alcohol (Figure 4). Wild-type mice preferred more alcohol than the DAT-Cnr2 mice during the 5-day test with acute stress (as determined by Student’s t-test comparison between genotypes). These results suggest that the DAT-Cnr2 KO mice are resistant to alcohol consumption even in acute stress conditions. This supports the previous report that CB2 receptor agonists is involved in the development and enhancement of alcohol preference in stressed but not in non-stressed control mice (Ishiguro et al., 2007; Onaivi et al., 2008).


Figure 4. Preference ratio for alcohol over water in normal conditions (A) and in response to an acute stress protocol (B) exhibited by wild-type mice (white circles) and DAT-Cnr2 cKO mice (black circles) during 5 consecutive days. Data presented as the Mean ± standard error of the mean (SEM) daily consumption ratio over 5 days. Significant difference between genotypes paired t-test (*P < 0.05, **P < 0.01, ***P < 0.001).


Figure 5. Deletion of CB2Rs in dopamine neurons (DAT-Cnr2 cKO) mice and alcohol preference. 8% alcohol-induced CPP (P < 0.05) in wild-type mice but the DAT-Cnr2 cKO mice did not show significant difference in post-conditioning phase between alcohol and saline.

Notes

Animal considerations:

  1. It is important to conduct preliminary studies to characterize the behavioral phenotype with particular equipment, environment, and animals. We measured baseline behaviors of wild-type, heterozygous, and homozygous of the DAT-Cre and DAT-Cnr2 mouse littermates to avoid genetic confounds. This is especially true in the case of varying mouse strains, either inbred or outbred, or if mice have been surgically or behaviorally manipulated prior to alcohol preference test.
  2. If required, the baseline of consumption can be established by weighing both bottles filled with 150 ml of water and weighed with the tops on and placed on top of the cages for three days.

Acknowledgments

This protocol is in memory of our co-author, Norman Schanz, whose sudden and tragic loss will be difficult to replace in the animal laboratory. This research was supported by William Paterson University. ACA was supported by the Mexican National Council of Science and technology (CONACYT # CVU332502/232728). HI was supported by Public Interest Trust Research Aid Fund for Stress-Related Diseases (with Commemoration of Imai Kimi), KAKENHI (18023009, 20023006, and 20390098). ESO was supported by NIH grant DA032890. QRL is supported by IRP/NIA/NIH. We thank the Dean, Dr. Venkat Sharma, of the College of Science and Health at William Paterson University for student worker support in maintaining our mice in the animal laboratory. We are also thankful for the technical assistant of Sneha Tammareddy, Eugene Dennis, Branden Sanabria, Paola Velandia, Steve Gross, and Monika Chung while working in the laboratory of ESO. This protocol and figures are adapted from our previous work (Liu et al., 2017).

Competing interests

The authors have no conflicts of interest to declare.

Ethics

The experimental procedures followed the Guide for the Care and Use of Laboratory Animals and were approved by William Paterson University Animal Care and Use Committee (IACUC).

References

  1. Barkley-Levenson, A. M. and Crabbe, J. C. (2012). Bridging animal and human models: Translating from (and to) animal genetics. Alcohol Res 34(3): 325-335.
  2. Cicero, T. (1980). Alcohol self-administration, tolerance and withdrawal in humans and animals: theoretical and methodological issues. In: Rigter, H. and Crabbe J. C. (Eds.) Alcohol tolerance and dependence. Amsterdam 7 Elsevier/North Holland Biomedical Press, pp 1-51.
  3. Crabbe, J. C., Phillips, T. J. and Belknap, J. K. (2010). The complexity of alcohol drinking: studies in rodent genetic models. Behav Genet 40(6): 737-750.
  4. Cunningham, C. L., Fidler, T. L. and Hill, K. G. (2000). Animal models of alcohol's motivational effects. Alcohol Res Health 24(2): 85-92.
  5. Ishiguro, H., Iwasaki, S., Teasenfitz, L., Higuchi, S., Horiuchi, Y., Saito, T., Arinami, T. and Onaivi, E. S. (2007). Involvement of cannabinoid CB2 receptor in alcohol preference in mice and alcoholism in humans. Pharmacogenomics J 7(6): 380-385.
  6. Liu, Q. R., Canseco-Alba, A., Zhang, H. Y., Tagliaferro, P., Chung, M., Dennis, E., Sanabria, B., Schanz, N., Escosteguy-Neto, J. C., Ishiguro, H., Lin, Z., Sgro, S., Leonard, C. M., Santos-Junior, J. G., Gardner, E. L., Egan, J. M., Lee, J. W., Xi, Z. X. and Onaivi, E. S. (2017). Cannabinoid type 2 receptors in dopamine neurons inhibits psychomotor behaviors, alters anxiety, depression and alcohol preference. Sci Rep 7(1): 17410.
  7. Mardones, J. and Segovia-Riquelme, N. (1983). Thirty-two years of selection of rats by ethanol preference: UChA and UChB strains. Neurobehav Toxicol Teratol 5(2): 171-178.
  8. Onaivi, E. S., Carpio, O., Ishiguro, H., Schanz, N., Uhl, G. R. and Benno, R. (2008). Behavioral effects of CB2 cannabinoid receptor activation and its influence on food and alcohol consumption. Ann N Y Acad Sci 1139: 426-433.
  9. Rhodes, J. S., Best, K., Belknap, J. K., Finn, D. A. and Crabbe, J. C. (2005). Evaluation of a simple model of ethanol drinking to intoxication in C57BL/6J mice. Physiol Behav 84(1): 53-63.
  10. Yang, X., Wang, S., Rice, K. C., Munro, C. A. and Wand, G. S. (2008). Restraint stress and ethanol consumption in two mouse strains. Alcohol Clin Exp Res 32(5): 840-852.
  11. Yoneyama, N., Crabbe, J. C., Ford, M. M., Murillo, A. and Finn, D. A. (2008). Voluntary ethanol consumption in 22 inbred mouse strains. Alcohol 42(3): 149-160.

简介

酒精偏好模型是与酒精中毒相关的最广泛使用的动物模型之一。压力源增加酒精消耗。在这里,我们提出了一个快速和有用的工具协议,以测试小鼠的酒精偏好和压力诱导的酒精消费。在该模型中,给动物两瓶,一瓶用乙醇在水中稀释的溶液,另一瓶用自来水。在几天内监测每瓶的消耗24小时,以评估动物对乙醇溶液相对于水的相对偏好。在第二阶段,通过每天约束它们一小时来对动物施加压力,并评估它们随后对自来水或乙醇溶液的偏好。优选通过每日消耗的体积和/或重量或液体来测量,然后将其转换为优选比率。将酒精偏好模型与条件性位置偏好范例相结合,以确定DAT- Cnr2 > Cre重组条件性敲除(cKO)小鼠中多巴胺能神经元中CB2大麻素受体缺失后的酒精调节和偏好。与野生型对照小鼠比较。

【背景】可以通过动物模型研究酒精中毒和饮酒的许多方面。酒精诱导积极的强化,动物可以寻求酒精,甚至为它工作。然而,酒精也可以是负面强化,因为它能够减少焦虑。没有动物模型能够复制酗酒的复杂特征。口服乙醇自我施用被广泛用于检查与理解酒精中毒相关的行为和生理学的特定方面(Mardones和Segovia-Riquelme,1983; Cunningham 等人,>,2000)。小鼠可以在细胞类型特异性水平上进行遗传操作,因此对于研究酒精中毒的细胞类型特异性遗传决定因素是有价值的。

酒精偏好模型是与酒精中毒相关的最广泛使用的动物模型之一。该模型符合重要标准,即乙醇应自口服(Cicero,1980; Crabbe et al。>,2010)。动物的基因型在该模型中对自我管理产生强烈影响。一些小鼠品系,如小鼠C57BL / 6J的近交品系(Rhodes et al。>,2005),对乙醇具有遗传影响的高度偏好,并且他们自愿口服它(Yoneyama et al 。>,2008; Barkley-Levenson和Crabbe,2012)。条件性地方范式(CPP)被广泛用于探索包括酒精在内的成瘾物质的影响,利用学习的联想。因此,酒精CPP测量酒精与特定环境的关联,以确定小鼠是否可以获得酒精CPP。

压力可以与正在进行的乙醇消耗相互作用以引发增加的摄入量(例如,>,自我治疗行为),从而增加对酒精使用障碍的初始易感性。在压力源中,急性和慢性压力的约束模型可以增加乙醇的消耗(Yang et al。>,2008)。

新的进展和积累的证据支持内源性大麻素系统在酒精影响方面的作用。内源性大麻素系统由两种大麻素受体CB1Rs和CB2Rs组成,具有内源性大麻素和用于生物合成和内源性大麻素失活的酶。我们的目标是总结用于测量酒精偏好的方案以及应激诱导的酒精消耗。我们还提供证据表明,在小鼠模型中多巴胺能神经元特异性缺失CB2R后,内源性大麻素系统在酒精偏好中发挥作用(Liu et al。>,2017)。

关键字:酒精, 应激, 小鼠模型, 行为, 大麻素, 大麻素位置偏爱

材料和试剂

  1. 50毫升带附帽的聚丙烯离心管(Boekel Scientific,目录号:120021)
  2. 带吸管盖的瓶子(见图1B和1C)(耐嚼,目录号:101445)
  3. 小鼠:成年(7周龄或更大)小鼠(C57BL / 6J)(THE JACKSON LABORATORY,目录号:000664)
    注意:也可以使用小鼠的替代菌株和年龄。将小鼠单独饲养,各自置于其各自的笼中,在温度受控(约23℃)和湿度12-12小时光 - 暗循环的环境中,可自由获取食物。有关详细信息,请参阅Notes中的动物注意事项。>
  4. 100%酒精和稀释液:8%,16%和32%(Sigma-Aldrich,目录号:1012768)


    图1.显示用于评估酒精偏好的实验装置的照片。 A.幼稚或条件性敲除小鼠和野生型对照受到急性应激的管。 B.管,水瓶和干净的空鼠笼。 C.对照笼的图像。 D.用于条件性位置偏好的装置的图像。

设备

  1. 鼠标聚碳酸酯家用笼(7.5 x W x 11 in L x 5 in H),带标准木片鼠标垫(Fisher Scientific,目录号:01-286-13A)
    制造商:Tecniplast,目录号:1290D00SU。>
    注意:在适应期间允许标准笼子更换(见下文)。但是,建议在数据收集期间避免更换笼子以防止泄漏。可以在阶段之间清洁笼子。每个主题鼠标一个笼子。>
  2. 不锈钢线笼盖改装,允许两瓶和食物的空间(Fisher Scientific,目录号:01-286-13A)
    制造商:Tecniplast,目录号:1290D00SU。>
  3. 用小(1毫米)钻头钻孔(在50毫升聚丙烯离心管顶部制作三个小孔,让鼠标呼吸,并在盖子上插一个较大的孔以插入尾部)
  4. Thermo Scientific称重秤(Thermo Fisher Scientific,目录号:20031)

软件

  1. Activity Monitor软件(Med Associates,St。Albans,VT)(用于自动数据收集)
  2. GraphPad PRISM 6.0软件(GraphPad Software.Inc。,San Diego,CA,USA)(用于数据分析)

程序

所有实验均在暗光阶段,在昏暗的光照下进行。

  1. 实验装置
    1. 动物:使用NaïveC57B1/ 6Js,并作为DAT- Cnr2 >条件性敲除(cKO)小鼠的野生型对照。先前报道了cKO小鼠的产生和繁殖(Liu 等人,>,2017)。
    2. 在受控的湿度和温度下,实验空间应安静且昏暗。
    3. 所有笼子应放在指定用于该实验的架子上。机架应稳固,以避免移动并防止泄漏。在取出瓶子时,实验者应该小心,以尽量减少泄漏量。应避免泄漏,以减少不希望的变化。
    4. 实验区域应清洁,所有测量应在一天的同一时间进行。
    5. 其中一个笼子将是“控制笼”。除了鼠标之外,这个笼子将具有上述所有内容。
    6. 所有笼子都应标有特定实验的信息,例如小鼠的品系,年龄和性别。食物应始终保持在右侧以保持一致性。

  2. 设置2瓶选择测试
    1. 用150毫升水填充所有瓶子并放在吸管上。
      注意:通过翻转瓶子确保没有漏水。>
    2. 用永久性标记中的主题鼠标(1,2,3 ......)标记每个瓶子。用相同的数字标记笼子。
    3. 将2个瓶子放在电线盖上(图2),尽量少摇动以避免滴水。两个瓶子应放在分开的金属架的一侧,另一侧留下食物。将食物颗粒放在电线盖的另一侧。
      注意:确保食物始终在右侧,以保持一致。使小鼠适应于测试室,具有与菌落室相同的条件(22±2℃;湿度:55±5%; 12小时暗淡/人造光/ 12小时黑暗循环:在下午7点关灯)。 >

  3. 习惯于2瓶
    1. 在受控条件下将一只小鼠放入适当的小鼠笼中(参见设备)(参见材料和试剂)。
    2. 将受试者分离成单独的笼子,并给它们两个装有150ml水的瓶子。在这些条件下让小鼠适应环境至少4天。

  4. 为酒精偏好设置2瓶选择
    1. 取下瓶子并清洁它们。
    2. 将水与100%酒精混合至16%。
    3. 对于每只小鼠,用150ml水填充一瓶,用酒精溶液填充另一瓶。将吸管盖放在瓶子上。
      注意:通过翻转瓶子确保没有漏水。>
    4. 用指定数量的小鼠标记每个瓶子,用永久性标记物(例如>,1 H 2 O / 1 EtOH)标记水或酒精。
    5. 记录瓶盖上的重量并记下记录表上的值。

  5. 评价酒精偏好
    1. 将2个瓶子放在电线盖上,轻轻摇动以避免滴水。两个瓶子应放在分开的金属架的一侧,另一侧留下食物。将食物颗粒放在电线盖的另一侧。
      注意:确保食物始终位于右侧,以保持一致。>
    2. 每天评估水和酒精溶液的消耗量,持续5天。记录重量。监测液体流动或溢出的任何问题,例如>,吸管下方的湿垫层。每天切换瓶子的位置(左侧与右侧)。
    3. “控制笼”只有两瓶没有鼠标的水。称量这些瓶子以确定倒置瓶子时的溢出量,并稍后校正其他组的数据。

  6. 急性应激诱导的酒精偏好(5天)
    1. 设置2瓶酒精选择(如上所述)。
    2. 在第一天取出瓶子进行每日称重后,将小鼠随机分配到实验组或对照组。用“控制”或“实验”标记笼子(在鼠标编号所在的位置上方)并将它们按组放在架子上。将实验组的小鼠约束在50ml锥形管中1小时。之后,他们被释放,并为两组(控制和实验)的瓶子再次放入笼子。
    3. 连续5天重复这些步骤。每天称量瓶子,并在每个阶段之前和之后称重小鼠。
    4. 酒精偏好的一般时间表由5天内的程序D-F描述。&nbsp;

  7. 慢性轻度压力(CMS)诱导饮酒(4-7周)(表1)
    1. 在单独的房间中为实验组和对照组设置单独的野生型和选择的cKO小鼠组(每组n = 6)。除非应激对照组外,所有动物均单独圈养。
    2. 对于CMS方案,按照半随机方案表1对小鼠施加各种应激物,按计划进行4-7周。
    3. 每周的压力方案包括食物剥夺12小时,缺水12小时,潮湿的床上用品12小时,隔夜频闪照明,尾部悬浮10分钟,管约束30分钟,如上所述,大声的音乐过夜,金属丝网取代床上用品12小时,30分钟将入侵者鼠标引入笼中并关闭或打开灯。这些压力因素在早晨或过夜配对。
    4. 独立房间内的所有非CMS组都随时获得食物和水。

  8. 条件性地方偏好(CPP)
    使用Med Associates(St. Albans,VT,USA)的红外光束探测器开放式野外装置(ENV-510)进行酒精CPP训练和测试。
    1. 使用来自Med Associates(St.Albans,VT,USA)的红外光子探测器开放式野外装置(ENV-510)进行酒精CPP训练和测试,该装置配备有两室位置偏好插入物(ENV-512),如如图1D所示。舱室1的地板具有平行杆(半径3毫米,中心间距8毫米),外面覆盖黑色纸板纸。隔室-2有一个不锈钢丝网(6 x 6)地板,白色纸板纸覆盖在墙壁外面。每个隔间为13厘米(宽)×15厘米(深)。
    2. 在三个阶段(图3)(预处理,调节和后处理阶段)中执行每组中的所有调节步骤和偏好测试。
    3. 在测试日的后调节阶段期间,允许小鼠探查两个隔室15分钟。
    4. CPP评分定义为在酒精配对隔室中花费的时间减去在CPP测试期间在盐水配对隔室中花费的时间。


      图2.设置2瓶选择酒精偏好和水可用


      图3.使用的CPP范例的图形表示。 所有调理会话和偏好测试每天在同一时间进行。在调节阶段之前使动物在CPP装置中适应45分钟。

    表1.慢性轻度压力因素(4-7周)&nbsp;

数据分析

所有数据都在电子表格中转录。为原始重量数据设置一列,并在其旁边显示已更正的重量数据列。校正后的重量数据是每天从对照笼中溢出(水和酒精)的量。该数字将从原始重量数据中减去。每天,必须从原水和酒精重量数据中减去溢出量,以确定校正后的重量。
&NBSP;为了计算每天消耗的酒精或水的量,每天平均瓶重。然后从当天每组的瓶重的平均值中减去这些值。从前一天减去新的瓶子重量,给小鼠一夜之间饮用的酒精或水量。
&NBSP;第0天是瓶子最初装满150毫升水/酒精,因此当天的消耗量是0.
&NBSP;酒精偏好率是通过将消耗的酒精量除以消耗的总液体(酒精+水)来确定的。
&NBSP;活动监视器使用光电断束计数记录在每个隔室中花费的时间。记录每只小鼠在隔室中花费的时间,并将CPP得分定义为在配对的隔室中花费的时间减去在CPP测试期间在盐水配对隔室中花费的时间。活动监控软件(Med Associates,St。Albans,VT)用于自动数据收集。

代表性数据:图4是酒精偏好测试的代表性数据。在该试验中,将小鼠单独饲养并用两个可用于三天的流体瓶进行习惯。我们使用两组:12只雄性C57 / 6J小鼠作为野生型和12只雄性DAT- Cnr2 >条件性敲除(cKO)小鼠,它们在中脑多巴胺神经元中不表达CB2R。在实验当天,将所有动物称重。一瓶装满150毫升自来水,另一瓶装150毫升16%乙醇溶液。将它们与顶部称重并放置在笼子顶部(正确标记)。对于酒精偏好测试,在上午10点对每只动物称重瓶子连续五天。不同笼子中瓶子的位置随机放置在笼子的哪一侧。在实验的第二部分,然后将每组分成两组。一组(n = 6)通过将它们放入50ml管中每天1小时而连续5天受到压力。剩下的动物留在笼子里。每个动物的所有瓶子同时连续五天称重,并且所有瓶子在上午11点放置在笼子的顶部。在所有实验中,计算消耗的醇与水的比率和总流体消耗量以获得优选比率。通过将消耗的总液体(酒精+水)消耗的酒精量除以亚急性应激来确定酒精偏好比。野生型小鼠优选在水上的醇,通过显着增加(通过重复测量ANOVA测定)显而易见。 DAT- Cnr2 > cKO小鼠(小鼠未显示酒精-CPP诱导,图5),并且未显示对酒精偏好的显着增加(图4)。在用急性应激进行的5天试验期间,野生型小鼠比DAT- Cnr2 >小鼠更喜欢酒精(通过Student's t > - 基因型之间的比较测定)。这些结果表明,即使在急性应激条件下,DAT- Cnr2 > KO小鼠也能抵抗酒精消耗。这支持了先前的报道,CB2受体激动剂参与了应激但不应用于非应激对照小鼠的酒精偏好的发展和增强(Ishiguro et al。>,2007; Onaivi et al 。>,2008)。


图4.野生型小鼠(白色圆圈)和DAT- Cnr2 >在正常条件下(A)和响应急性应激方案(B)对酒精对水的偏好比cKO小鼠(黑眼圈)连续5天。数据表示为5天内平均值(SEM)每日消耗比的平均值±标准误差。 t > - 测试的基因型之间存在显着差异(* P > <0.05,** P > <0.01,*** P >&lt; 0.001)。


图5.删除多巴胺神经元(DAT- Cnr2 > cKO)小鼠和酒精偏好中的CB2R。 8%酒精诱导的CPP( P >&lt; ; 0.05)在野生型小鼠中,但DAT- Cnr2 > cKO小鼠在酒精和盐水之间的调理后阶段没有显示出显着差异。

笔记

动物考虑:

  1. 重要的是进行初步研究以表征特定设备,环境和动物的行为表型。我们测量了DAT- Cre >和DAT- Cnr2 >小鼠同窝小鼠的野生型,杂合子和纯合子的基线行为,以避免遗传混淆。对于不同的近交或远交的小鼠品系,或者如果在酒精偏好测试之前对小鼠进行手术或行为操作,尤其如此。
  2. 如果需要,可以通过称量装有150ml水的瓶子并称重顶部并放置在笼子顶部三天来确定消耗基线。

致谢

该协议是为了纪念我们的共同作者Norman Schanz,他的突然和悲剧性的损失将很难在动物实验室取代。这项研究得到了威廉帕特森大学的支持。 ACA得到了墨西哥国家科学技术委员会的支持(CONACYT#CVU332502 / 232728)。 HI得到公共利益信托研究援助基金的应激相关疾病(纪念今井基米),KAKENHI(18023009,20023006和20390098)的支持。 ESO得到NIH资助DA032890的支持。 QRL由IRP / NIA / NIH支持。我们感谢威廉帕特森大学科学与健康学院的院长Venkat Sharma博士,感谢学生们在动物实验室维护我们的老鼠。我们还感谢Sneha Tammareddy,Eugene Dennis,Branden Sanabria,Paola Velandia,Steve Gross和Monika Chung的技术助理,他们在ESO实验室工作。该协议和数据改编自我们以前的工作(Liu et al。>,2017)。

利益争夺

作者没有利益冲突申报。

伦理

实验程序遵循实验动物护理和使用指南,并由William Paterson大学动物护理和使用委员会(IACUC)批准。

参考

  1. Barkley-Levenson,A。M.和Crabbe,J。C.(2012)。 桥接动物和人类模型:从(和)动物遗传学翻译。 酒精含量> 34(3):325-335。
  2. Cicero,T。(1980)。人类和动物的酒精自我管理,耐受和戒断:理论和方法问题。在:Rigter,H。和Crabbe J. C.(编辑)酒精耐受和依赖。>阿姆斯特丹7 Elsevier /北荷兰生物医学出版社,第1-51页。
  3. Crabbe,J.C。,Phillips,T.J。和Belknap,J.K。(2010)。 饮酒的复杂性:啮齿动物遗传模型的研究。 Behav Genet > 40(6):737-750。
  4. Cunningham,C。L.,Fidler,T。L.和Hill,K.G。(2000)。 酒精激励效果的动物模型。 酒精健康> 24(2):85-92。
  5. Ishiguro,H.,Iwasaki,S.,Teasenfitz,L.,Higuchi,S.,Horiuchi,Y.,Saito,T.,Arinami,T。和Onaivi,E。S.(2007)。 大麻素CB2受体参与小鼠酒精偏好和人类酒精中毒。 药物基因组学J > 7(6):380-385。
  6. Liu,QR,Canseco-Alba,A.,Zhang,HY,Tagliaferro,P.,Chung,M.,Dennis,E.,Sanabria,B.,Schanz,N.,Escosteguy-Neto,JC,Ishiguro,H。 ,Lin,Z.,Sgro,S.,Leonard,CM,Santos-Junior,JG,Gardner,EL,Egan,JM,Lee,JW,Xi,ZX和Onaivi,ES(2017)。 多巴胺神经元中的大麻素2型受体抑制精神运动行为,改变焦虑,抑郁和酒精偏好。 Sci Rep > 7(1):17410。
  7. Mardones,J。和Segovia-Riquelme,N。(1983)。 通过乙醇偏好选择大鼠32年:UChA和UChB菌株。 Neurobehav Toxicol Teratol > 5(2):171-178。
  8. Onaivi,E.S.,Carpio,O.,Ishiguro,H.,Schanz,N.,Uhl,G.R。和Benno,R。(2008)。 CB2大麻素受体激活的行为影响及其对食物和酒精消费的影响。 Ann NY Acad Sci > 1139:426-433。
  9. Rhodes,J。S.,Best,K.,Belknap,J.K.,Finn,D.A。和Crabbe,J.C。(2005)。 评估C57BL / 6J小鼠中毒饮酒的简单模型。 Physiol Behav > 84(1):53-63。
  10. Yang,X.,Wang,S.,Rice,K.C.,Munro,C.A。和Wand,G。S.(2008)。 限制两种小鼠品系中的压力和乙醇消耗量。 Alcohol Clin Exp Res > 32(5):840-852。
  11. Yoneyama,N.,Crabbe,J。C.,Ford,M。M.,Murillo,A。和Finn,D。A.(2008)。 22种近交小鼠品系的自愿乙醇消耗量。 酒精> 42(3):149-160。
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引用:Canseco-Alba, A., Schanz, N., Ishiguro, H., Liu, Q. and Onaivi, E. S. (2018). Behavioral Evaluation of Seeking and Preference of Alcohol in Mice Subjected to Stress. Bio-protocol 8(20): e3061. DOI: 10.21769/BioProtoc.3061.
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