We thank Dr. J-L. Galzi (UMR7242 CNRS; Illkirch, France) for his support. 
This work was supported by the CNRS, INSERM, Universit&#233; de Strasbourg, Alsace BioValley and by grants from Conectus, Agence National de la Recherche (ANR 08 EBIO 014.02) Conseil R&#233;gional d&#8217;Alsace (Pharmadol), Communaut&#233; Urbaine de Strasbourg (Pharmadol), ICFRC (Pharmadol), OSEO (Pharmadol), Direction G&#233;n&#233;rale des Entreprises (Pharmadol).

All experiments were carried out in strict accordance with the European guidelines for the care of laboratory animals (European Communities Council Directive 86/609/ECC) and the ethical guidelines for investigation of experimental pain in conscious animals13. Male C57BL6/N Tac mice (10 weeks, 25 - 30 g) were housed in an institutional animal facility with a care staff in charge of operating the facility in compliance with environmental standards. Animals were housed in groups (maximum five mice per cage) under a 12 hr / 12 hr light / dark cycle at a constant temperature (21 &#177; 1 &#176;C) with free access to food and water. All experiments were performed at the same period of the day (10:00 am to 4:00 pm) using a cohort of 16 mice. Specific materials and equipment are indicated in the Materials table.
A five-step procedure to monitor morphine induced-hyperalgesia and tolerance
The protocol is divided into five sequential steps (A-E) over a duration of 15 days Figure 1.
1. Mice Handling (Step A; d-7 to d-5)
<ol>
	<li>Handle mice and habituate them to freely enter into the restrainer. This preliminary step reduces stress - thus minimizing any confusion with stress-induced analgesia - and allows the animals to be accustomed to the investigator, handling and manipulation into the mouse restrainer. Each mouse is gently handled for 5 min every day.</li>
</ol>
2. Basal Nociceptive Response (Step B; d-4 to d-1)
<ol>
	<li>Measure tail withdrawal latencies using the Tail Immersion Test (TIT):
	<ol>
		<li>Set the thermostat to 48 &#176;C.</li>
		<li>Gently introduce the mouse into the restrainer. Dip the protruding 2/3 end of its tail into the water bath and start the chronometer.</li>
		<li>Stop the chronometer as soon as the mouse withdraws its tail from the hot water and record the latency time (in sec). In the absence of any nociceptive reaction, a 25 sec cut-off is used to prevent tissue damage.</li>
		<li>Replace the mouse in its cage and test the next one until the end of the series.</li>
		<li>Repeat the nociceptive response measurements two more times, taking measurements from the animals in the same order. The nociceptive response latency (sec) for each mouse is determined as the mean value for three successive determinations.</li>
	</ol>
	</li>
	<li>Measure mechanical responses using the Tail Pressure Test (TPT)
	<ol>
		<li>Gently introduce the mouse into the restrainer and position its tail under the conic tip of the analgesimeter.</li>
		<li>Press the foot-switch to apply uniformly increasing pressure onto the proximal part of the tail until the first nociceptive reaction (struggling, squeaking) occurs. At the moment the animal reacts, record the current force (in grams) that elicits the nociceptive response. In the absence of any reaction, a 600 g cut-off value is used to avoid tissue damage.</li>
		<li>Repeat this measure on the median and distal parts of the tail of the same mouse. An interval of at least 30 sec is observed between measures on a given mouse to avoid adaptation or stress bias. Replace the mouse in its cage and test the next animal until the end of the series (i.e. all mice are tested). The nociceptive value (grams) for each mouse is taken as the mean value for three measurements (i.e. proximal, median and distal parts of the tail of each animal).</li>
	</ol>
	</li>
	<li>Repeat nociceptive testing (all the procedures outlined under step 2) on subsequent days, d-3 to d-1.</li>
</ol>
3. Measurement of Morphine Analgesia (Step C; d0)
<ol>
	<li>Define the best combination of animals that allows the selection of two groups (n=8 per group) of mice with stable and comparable average nociceptive values, whichever nociceptive modality (TIT or TPT) is considered. This value will be taken as the basal nociceptive response of reference for future &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups.</li>
	<li>Measure the body weight of each animal.</li>
	<li>Prepare a morphine solution (0.5 mg morphine per ml) in sterile physiological saline (NaCl 0.9%) for subcutaneous administration (5 mg morphine per kg of animal&#8217;s body weight).</li>
	<li>Measure the nociceptive response latency (taken as time point 0) for each mouse of both &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups in the TIT (all steps under 2.1 above). Then measure nociception in the TPT (all steps under 2.2 above).</li>
	<li>Inject subcutaneously morphine (typically 0.25 ml of a 0.5 mg/ml morphine solution per 25 g mouse weight) and saline (0.25 ml per 25 g mouse weight) to the &#8216;morphine&#8217; and &#8216;saline&#8217; groups, respectively.</li>
	<li>Measure nociceptive values on the TIT and TPT (all procedures outlined above under steps 2.1 and 2.2, respectively) over a time-course (at 30 min interval) to assess morphine (5 mg/kg)-induced analgesia:
	<ol>
		<li>After 30 min post-injection, measure the nociceptive response (single determination) for each mouse of the &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups, using the TIT then the TPT.</li>
		<li>Then measure nociceptive response values (TIT and TPT) in all mice at time points (in hr): 1-1.5-2-2.5-3 and 3.5 post-injection.</li>
	</ol>
	</li>
</ol>
4. Chronic Morphine Treatment - Morphine-induced Hyperalgesia (Step D; d1 to d6)
<ol>
	<li>On Day: d1
	<ol>
		<li>Measure nociceptive response values on the TIT and TPT as described above (steps 2.1 and 2.2). Carefully annotate withdrawal latencies and pressure limits for each animal.</li>
		<li>Prepare a fresh morphine solution as detailed in step 3.2.</li>
		<li>Inject subcutaneously morphine (5 mg/kg b.w.) to the entire &#8216;Morphine&#8217; group and physiological saline (0.25 ml per 25 g mouse weight) to the &#8216;Saline&#8217; group. Let the animals rest until the next day.</li>
	</ol>
	</li>
	<li>On Days: d2, d3, d4, d5 and d6 repeat the operations described under Section 4.1</li>
</ol>
5. Evidence for Analgesic Tolerance (Step E; d7)
<ol>
	<li>Evaluate morphine-induced analgesia according to the time-course paradigm already detailed in Section 3.</li>
</ol>
6. Data Acquisition and Statistical Analyses
<ol>
	<li>Assessment of Basal Nociceptive Response Values (Step B)
	<ol>
		<li>Calculate for each day (over the d-4 to d-1 period) the mean &#177; SEM values (n=8) for basal nociceptive responses as afforded from TIT and TPT within the &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups.</li>
		<li>Plot mean basal nociceptive values as a function of time (day) for both groups Figure 2.</li>
	</ol>
	</li>
	<li>Analysis of Morphine Analgesia Time-course at Days d0 (Step C) and d7 (Step E)
	<ol>
		<li>Calculate, at each time point after morphine injection, the mean &#177; SEM values (n=8) for nociceptive responses as afforded from TIT (in sec) and TPT (in g) in each group.</li>
		<li>Plot mean nociceptive response values as a function of time for the &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups at day 0 Figure 3 and day 7 Figure 5.</li>
	</ol>
	</li>
	<li>Development of Morphine-induced Hyperalgesia (Step D)
	<ol>
		<li>Calculate for each day (over the d0-d7 treatment period) the mean &#177; SEM values (n=8) for basal nociceptive response values as afforded from TIT and TPT within the &#8216;Saline-treated&#8217; and &#8216;Morphine-treated&#8217; groups.</li>
		<li>Plot mean basal nociceptive response values as a function of time (day) for the &#8216;Saline-treated&#8217; and &#8216;Morphine-treated&#8217; groups Figure 4.</li>
	</ol>
	</li>
	<li>Evidence for Analgesic Tolerance (Steps C and E)
	<ol>
		<li>Determine from the morphine time-course experiment performed at d0 Figure 3 the time value (or time range) required for morphine to induce a maximal analgesic response.</li>
		<li>Take this value (usually 30 min) as the reference time to estimate at d7 Figure 5 the nociception baseline value (saline-treated group) and the actual analgesic response (morphine-treated group) to acute morphine.</li>
		<li>Nociceptive values taken at time point 30 min from morphine time-course experiments performed at d0 and d7 for the saline-treated and morphine-treated groups, are presented as histograms Figure 6.</li>
		<li>Statistics: Analyze data using one-way repeated measures ANOVA. The factors of variation were treatment (between subjects) and time (within subject). To check differences separately in each group, repeated measures ANOVA was performed. Comparisons between two groups were performed using unpaired t-test or paired t-test when appropriate.</li>
	</ol>
	</li>
	<li>The Level of Significance is Set at P&#60;0.05. All Statistical Analyses are Carried out using the STATview Software.</li>
</ol>

The authors declare that they have no competing financial interests.

Critical Steps
Choice of the animal model for nociception measurements
Variability in nociceptive and analgesic sensitivity among mice strains has been examined (reviews&#160;14-16)&#160;reviews using various pain models differing in their etiology (nociceptive, inflammatory, neuropathic), modality (thermal, chemical, mechanical), duration (acute, tonic, chronic) and site of administration (cutaneous, subcutaneous, visceral). When compared to other strains, C57BL/6J (&#8220;J&#8221; for Jackson Laboratory) mice became a popular animal model for pain studies as they exhibit a high basal nociceptive sensitivity17,18 and a moderate analgesic response to opiates14,19. Following chronic morphine treatment, they also develop significant analgesic tolerance20,21, hyperalgesia21,22 and dependence20,23.
Here, experiments were performed on C57BL/6N Tac mice (&#8220;N&#8221; for National Institute of Health and &#8220;Tac&#8217; for Taconics farm) which belong to a separate branch of the B6 lineage. Although C57BL/6 mice have been long considered as interchangeable, recent studies pointed to significant behavioral differences among C57BL/6J and C57BL/6N strains24. In particular, the lower sensitivity of the three C57BL/6N substrains (including the Tac one) to acute thermal pain may be regarded as an advantage for testing this phenotype.
Male mice were selected as the vast majority of pain studies, using mice as the animal model, are performed on juvenile males25. In our hands, they provided robust and reproducible data when examined from the analgesia or hyperalgesia points of view. Occasionally, we noticed a tendency for C57BL/6N females to provide more variable responses, both in the TIT and TPT tests. Although this observation may reflect natural variations linked to the hormonal status of females, overall mechanisms underlying sex differences in pain and analgesia still remain a matter of controversy. Some aspects of this hot debate will be briefly presented in the next &#8216;Limitations of the technique&#8217; section.
Animal habituation
Mice were first allowed to get accustomed to the animal facility during one week. Similar to any other behavioral study, testing was performed following a 3 day-acclimatization period (Figure 1, step A). As nociceptive tests are sensitive to stress, first measures may give longer latencies than subsequent ones, especially in non-habituated mice26,27. The habituation step allows also the obtention of more stable nociceptive response values within the same day and between days Figures 2 and&#160;4. To reduce circadian effects on nociceptive and analgesic sensitivity28,29, all testings were conducted between 10:00 am and 4:00 pm.
Selection of nociceptive tests
Nociceptive tests use either thermal, mechanical, chemical or electrical stimuli (review&#160;26,27,30. Their choice is critical as different nociceptive modalities may be processed through different nociceptors and fibers18,31,32.
We selected the tail immersion test (TIT)33, a modified version of the classical tail flick test developed by D&#8217;amour and Smith34, and the tail pressure test (TPT), adapted from Randall and Selitto35, as examples of thermal and mechanical modalities to study morphine-induced analgesia, hyperalgesia and tolerance in mice. Both tests have been widely used in rats. A cut off time was systematically defined to avoid or limit the risk of tissue damage.
Morphine-induced analgesia, hyperalgesia and tolerance
Morphine, the prototypical mu-opiate agonist, was selected here as it is a potent analgesic and OIH-inducer, both in humans and mice1,2,36. Morphine analgesic potency is known to vary with mice strains, routes of administration and nociceptive modalities. In C57BL/6 mice, reliable analgesia is usually obtained following subcutaneous injections of morphine in the 1-20 mg/kg dose range14,21. Accordingly, we chose to study acute analgesia following a single administration (s.c.) of morphine at 5 mg/kg, close to its ED50 value (7-20 mg/kg) assessed from thermal nociception19,21.
Repeated morphine administration is often accompanied with analgesic tolerance (evidenced either from a rightward shift of the dose-response curve or from a decrease in analgesic response amplitude or duration) and hyperalgesia (an exacerbated sensitivity to painful stimuli evidenced from a decrease in basal nociceptive value). Both adverse phenomena depend on rodent strains, on the nature of the opiate compound which is selected and its dosage, on treatment duration and on nociceptive modalities21. For example, experimental paradigms to study tolerance and hyperalgesia consist in daily administration of high and constant (20 to 40 mg/kg per day)22 or of escalating (up to 50 or even 200 mg/kg)20,21 morphine doses. Accordingly, we promoted the development of hyperalgesia and tolerance in C57BL/6 mice through daily morphine administration (5 mg/kg; s.c.) over a 8-day period. This moderate morphine dose was preferred over higher ones to better mimic clinic usage.
Set up of TIT operational window
A possible pitfall in TIT might be related to the role of the tail in the thermoregulation of rodents26,37. As ambient temperature is a key factor in nociceptive response variations, it should be kept constant (here at 21 &#176;C) throughout experiments38. Heat intensity is usually set up to detect a nociceptive response within 5 to 10 sec27. Indeed, greater latencies may increase the risk for monitoring animal movements unrelated to the nociceptive stimulus, whereas shorter ones may reduce the differential power of the test. We performed TIT measurements at a fixed temperature of 48 &#176;C. Tail withdrawal latencies were close to 9 sec (basal nociceptive value) and varied from 4 sec (hyperalgesia) to 25 sec (maximal analgesia; cut off). In addition to practical reasons, measurements of nociceptive response values at a fixed temperature may a priori involve the same repertoire of nociceptors and circuits, thereby facilitating data interpretation.
Possible Modifications 
Optimization of the TIT operational window for analgesia and OIH measurements
When focusing on an analgesic response, low baseline values (higher heat intensity) may favor the detection of a delay in the response. In turn, to address the consequence of a painful stimulus or the development of OIH, higher baseline values (lower heat intensity; here 48 &#176;C) may facilitate the detection of faster responses Figure 4.
Although we found morphine at 5 mg/kg a convenient dose to induce a robust analgesic response Figure 3&#160;and to promote (upon repeated administration) significant hyperalgesia Figure 4, its dosage may be adapted as mentioned before (Critical step : Morphine-induced analgesia, hyperalgesia and tolerance). For example, lower doses may be used to reduce analgesia amplitude (thereby avoiding cut-off limitations) whereas higher doses may be chosen to accelerate hyperalgesia onset and increase its amplitude.
Overall, optimization of the &#8216;nociceptive window&#8217; should be adapted to the genetic background of mice under study and take into account the possibility for the involvement of distinct arrays of nociceptors and circuits.
Alternative opiate agonists (fentanyl, remifentanyl)
Although most clinically used opiates target the mu-opioid receptor as agonists, they differ considerably with respect to their pharmacological properties both in vitro and in vivo. For example, remifentanyl and fentanyl, in marked contrast with morphine, behave as full agonists and promote internalization of mu-opioid receptors39. Opiate analgesics such as morphine and fentanyl have half-lives in the range of hours40, while remifentanyl has an ultra-short half life of several minutes41. In humans, best evidence for OIH is from patients who received opiates during surgery, including short-acting compounds such as remifentanyl2,42. Thus, fentanyl and remifentanyl may be valuable tools too to study the development of hyperalgesia and tolerance in mice, under TIT and TPT paradigms.
Alternative modes of induction of OIH (chronic vs acute administration)
OIH is seen in humans and animal models as a consequence of opiate administration, whether at very low or extremely high dosages1,2. We report here on OIH development following chronic treatment of mice with moderate doses of morphine. Several days of treatment of C57BL/6N mice were necessary to evidence a clear and reproducible hyperalgesic state Figure 4. Daily morphine injections could be adequately replaced with implanted morphine pellets : upon their removal, both thermal hyperalgesia and mechanical allodynia have been already reported in mice43. Infusion of opiates through a micro-osmotic pump is another possibility44. In rodents, long-lasting hyperalgesia is also achievable following acute administration of fentanyl using a protocol mimicking the use of this mu-opioid agonist in human surgery36,45,46.
Limitations of the Technique
Animal species and models for pain
Comparative studies of numerous mouse strains provided evidence for great variations in nociceptive responses to painful stimuli17,31,47 and in OIH levels following 4-days morphine treatment22. Whether mechanisms underlying pain processing and modulation in animal models (mice and rats) are relevant for chronic pain patients remains a fundamental and opened question. Thus, much caution should be paid to the interpretation of animal data and to their predictive validity for humans16.
Sex differences in pain and analgesia
Most preclinical studies on animal models for pain have been conducted on male rodents16,25,48. Despite this selection bias, the emerging view was to consider males as better responders to opiate analgesics49,50, less prone to develop opioid-induced hyperalgesia 51,52 and more tolerant to morphine analgesia53 than their female counterparts (review&#160;54). However, sex differences with regard to nociception and analgesic drugs efficacy do not resume in such &#8216;a one size fits all&#8217; paradigm. Indeed, a wealth of data now indicates that numerous variables may influence the magnitude and direction of sex differences such as opioid drug efficacy and selectivity, nociceptive assay, genetic background, age, gonado-hormonal status or social interaction48,54. In humans, clinical pain is more prevalent in women but whether this fact reflects actual sex differences remains a matter of debate48,55,56. For example, global analysis of fifty clinical trials indicated no significant differences in analgesic properties between genders whereas meta-analyses performed on patients-controlled subjects pointed to a significantly greater opioid efficacy in women57.The latter observation, which markedly contrasts with what has been found in rodents, again raises several questions regarding the origin for such divergences16,48,55,57. Altogether, sex differences in analgesia do exist and merit further focus on underlying mechanisms and clinical implications.
About nociceptive tests
The tail withdrawal test is a spinal reflex but it may be subject to supraspinal influences58. TIT is relatively easy to perform on rats but requires more expertise in mice. A potential difficulty is to maintain the mouse in a correct posture without inducing unwanted stress. The proposed protocol may be adjusted according to cohort size. 16 animals (8 control and 8 treated) are easily managed as far as measurement of their basal nociceptive response values (using TIT first, then TPT for the whole series of mice) is under concern. Monitoring analgesia time courses requires the establishment of a precise time schedule and the evaluation of the maximal number of animals that may be tested (TIT first, then TPT) within the imparted time interval (here 30 min). The whole cohort of animals may thus be divided into subgroups to allow the experimenter to respect kinetic limitations.
Significance of the Technique with respect to Existing / Alternative Methods 
OIH in rats versus mouse
Rats have been extensively used to study opioid analgesia, hyperalgesia and tolerance, following acute or chronic opiate administration46,59-61. Indeed, for several practical reasons, they may be considered superior to mice as an animal model for pain experiments16,61. However, until recently, the generation of genetically modified rats was not a straightforward procedure. As numerous genetically modified mouse strains are already available, our model offers the opportunity to study the contribution of numerous individual genes in OIH and tolerance development in mice.
TIT and TPT versus other nociceptive tests
TIT is a variant of the tail flick test, the most obvious difference being the area of stimulation. In contrast with radiant heat, immersion of the tail into hot water leads to a quick and uniform increase in its temperature. Compared to other forms of thermal nociception testing (hot-plate or Hargreaves tests), TIT provides fairly reproducible results both across and within subjects.
TPT is a very popular test for the study of mechanical nociception26,27,35 which probably involves distinct nociceptive fibers and molecular transducers then TIT32. It provides quick and reliable measurements59 but requires some expertise from the experimenter and large animal cohorts. As an alternative to the analgesimeter used in the present study, other procedures or apparatus relying on strain gauges do exist (review&#160;27). TPT is best suited for studying mechanical hyperalgesia whereas von Frey filaments are usually taken to evaluate mechanical allodynia (review&#160;27).
Future Applications or Directions after Mastering this Technique
The experimental OIH/tolerance model we present here may be similarly adapted to genetically modified mice in order to evaluate the role of individual genes in the modulation of pain. It also provides a model system to investigate the effectiveness of potential therapeutic agents to relieve chronic pain.

Opioid-induced hyperalgesia (OIH) and analgesic tolerance limit the clinical efficacy of opiates in animals and humans1-3. The involvement of pro-inflammatory4,5 or of pro-nociceptive (anti-opioid)6,7 systems are currently explored hypotheses. The elucidation of the mechanisms underlying OIH and tolerance necessitates a combination of in vivo and in vitro approaches, using appropriate animal models, experimental protocols and molecular tools.
Behavioral pharmacology is the dominant paradigm to monitor and quantify analgesic and hyperalgesic states in laboratory animals (rats, mice). The application of a noxious stimulus (thermal, mechanical or chemical) to a convenient body part (hindpaw, tail) of the animal leads to a nocifensive withdrawal that can be easily scored.
We propose here a methodological approach for inducing, recording and quantifying OIH and tolerance in wild-type mice, using the tail-immersion and tail pressure tests. The procedure allows an easy, sensitive and reproducible determination of thermal and mechanical nociceptive response values in mice. As demonstrated in the video protocol, C57BL/6 mice experience significant hyperalgesia following chronic morphine administration and maintain this for several days. Both thermal and mechanical nociceptive values are significantly reduced, compared to baseline measurements on na&#239;ve animals. Moreover, our experimental set-up allows to monitor, in addition to the development of OIH, the decline of the analgesic response to morphine (tolerance). Presented data support the view that hyperalgesia and tolerance may involve common cellular and molecular mechanisms8,9, although this is disputed in the literature1,10-12. Finally, this protocol may be similarly adapted to genetically modified mice in order to evaluate the role of individual genes in the modulation of pain. It also provides a model system to evaluate the effectiveness of potential therapeutic agents to improve opiate analgesic effects.

<table border="0" cellpadding="0" cellspacing="0" width="879">
	<tbody>
		<tr height="17">
			<td height="34" rowspan="2" style="height:34px;width:220px;">Name</td>
			<td rowspan="2" style="width:231px;">Company</td>
			<td rowspan="2" style="width:139px;">Catalog Number</td>
			<td rowspan="2" style="width:291px;">Comments/description</td>
		</tr>
		<tr height="17">
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;width:220px;">C57BL/6N Tac mice</td>
			<td style="width:231px;">Taconic, Ry, Denmark</td>
			<td style="width:139px;">C57BL/6N Tac B6-M</td>
			<td style="width:291px;">Male mice (25-30g)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">Morphine hydrochloride</td>
			<td style="width:231px;">Francopia, Paris, France</td>
			<td style="width:139px;">CAS n&#176; 52-26-6</td>
			<td style="width:291px;">Delivered with special authorization</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">Syringes (Terumo)</td>
			<td style="width:231px;">Dutscher, Brumath, France</td>
			<td style="width:139px;">050000</td>
			<td style="width:291px;">Polypropylene; Sterile; Volume : 1 ml</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">Needles (Terumo)</td>
			<td style="width:231px;">Dutscher, Brumath, France</td>
			<td style="width:139px;">050101</td>
			<td style="width:291px;">G26 &#189; (Terumo reference : NN2613RO1)</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:220px;">Mouse restrainer</td>
			<td style="width:231px;">Home-made</td>
			<td style="width:139px;">-</td>
			<td style="width:291px;">Two metallic grids (5x11cm) assembled with adhesive tape and staples.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">Thermostated water bath GR150</td>
			<td style="width:231px;">Grant Instruments, Cambridge, UK</td>
			<td style="width:139px;">GP 0540003</td>
			<td style="width:291px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">Analgesimeter</td>
			<td style="width:231px;">Panlab, Barcelona, Spain</td>
			<td style="width:139px;">LE 7306</td>
			<td style="width:291px;"></td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;width:220px;">Kaleidagraph software</td>
			<td style="width:231px;">Synergy software, Reading, PA, USA</td>
			<td style="width:139px;">Kaleidagraph 4.03&#160;</td>
			<td style="width:291px;">Scientific graphing</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:220px;">STATview software</td>
			<td style="width:231px;"></td>
			<td style="width:139px;"></td>
			<td style="width:291px;">Free download; Statistics</td>
		</tr>
	</tbody>
</table>

assessment of morphine-induced hyperalgesia and analgesic tolerance in mice using thermal and mechanical nociceptive modalities

Opioid-induced hyperalgesia and tolerance severely impact the clinical efficacy of opiates as pain relievers in animals and humans. The molecular mechanisms underlying both phenomena are not well understood and their elucidation should benefit from the study of animal models and from the design of appropriate experimental protocols. 
We describe here a methodological approach for inducing, recording and quantifying morphine-induced hyperalgesia as well as for evidencing analgesic tolerance, using the tail-immersion and tail pressure tests in wild-type mice. As shown in the video, the protocol is divided into five sequential steps. Handling and habituation phases allow a safe determination of the basal nociceptive response of the animals. Chronic morphine administration induces significant hyperalgesia as shown by an increase in both thermal and mechanical sensitivity, whereas the comparison of analgesia time-courses after acute or repeated morphine treatment clearly indicates the development of tolerance manifested by a decline in analgesic response amplitude. This protocol may be similarly adapted to genetically modified mice in order to evaluate the role of individual genes in the modulation of nociception and morphine analgesia. It also provides a model system to investigate the effectiveness of potential therapeutic agents to improve opiate analgesic efficacy.

Assessment of Basal Nociceptive Values of Na&#239;ve Mice (Step B)
TIT and TPT were sequentially applied to the whole cohort of mice (n=16), providing mean nociceptive response values. Best combination of animals allowed an a posteriori definition of two groups (n=8) of mice, referred to as Saline and Morphine, which display similar and stable basal nociceptive values Figure 2. The equivalence of both groups is valid whatever the nociceptive test (TIT: Figure 2A; TPT : Figure 2B) that was selected.
Time-course for Morphine Analgesia at Day 0 (Step C)
Morphine analgesia was evaluated following a single injection (s.c.) of morphine (5 mg/kg) in na&#239;ve mice using both TIT Figure 3A and TPT Figure 3B. In both tests, statistical analyses with one way repeated measures ANOVA reveal that there is a significant interaction between treatment and time for TIT (F(7, 98) = 72, p&#60;0.001) and TPT (F(7, 98) = 31, p&#60;0.001). TIT and TPT data analyses using repeated measures ANOVA indicate that there is no effect of saline injection (F(7, 49) = 0.49, p&#62;0.05) and F(7, 49) = 1.85, p&#62;0.05 respectively for TIT and TPT tests), whereas morphine injection induces a strong analgesia in mice (F (7, 49) = 92.46, p&#60;0.001) and F(7, 49) =34.37, p&#60;0.001 respectively for TIT and TPT tests). The maximal analgesic effect of morphine was reached after 30 min in TIT and after 60 min in TPT as compared to saline-injected controls (p&#60;0.001, unpaired t-test).
Repeated Morphine Administrations Induces Hyperalgesia in Mice (Step D)
Basal nociceptive values were measured every day before saline or morphine administration (see protocol). As shown in Figure 4, once daily morphine administrations over a 6 days treatment period induced a significant and progressive lowering of thermal (F(7, 56) = 11.6, p&#60;0.001, repeated measures ANOVA; Figure 4A) and mechanical (F(7,56) = 15,55, p&#60;0.001, repeated measures ANOVA; Figure 4B) basal nociceptive values. Hyperalgesia developed rapidly as it started to be significant at day 1 in TIT (p&#60;0.01, unpaired t-test, compared to saline-injected controls) and at day 2 in TPT (p&#60;0.05, unpaired t-test, compared to saline-injected controls).
Time-course for Morphine Analgesia at Day 7, after Chronic Morphine Treatment (Step E)
On day 7, mice that received daily morphine or saline injections over a 7-day period (d0 to d6) were examined in TIT Figure 5A and TPT Figure 5B first for their basal nociceptive values and then for their analgesic response to acute morphine (5 mg/kg, sc.). In agreement with the development of hyperalgesia shown in Figure 4, the basal nociceptive value (time 0) of mice that were chronically treated with morphine was significantly lower than that of saline-injected control mice (p&#60;0.001, unpaired t-test). Following acute morphine, the nociceptive response of the chronic morphine-treated group significantly increased, but only slightly exceeded the basal nociceptive value of saline-injected control mice measured at 30 min in TIT and TPT (p&#60;0.01 and p&#60;0.05, unpaired t-test, respectively) and at 60 min in TIT (p&#60;0.05; unpaired t-test). From 2 hr after morphine treatment until the end of experiment, nociceptive responses returned to lower values than those of control mice (p&#60;0.001, unpaired t-test).
Comparison of Maximum Analgesic Responses of Mice to Morphine before (Day 0) and after Chronic Morphine Treatment (Day 7).
Nociceptive threshold values presented in Figure 6 are from TIT (A) and TPT (B) performed 30 min after saline or morphine injection as illustrated in Figures 3 (day 0) and 5 (day 7). A strong decrease in morphine analgesia was observed in mice after chronic morphine treatment for 7 days in comparison with their initial analgesia response at day 0 in both nociceptive tests (p&#60;0.001 paired t-test). These data demonstrate that tolerance did develop in pain hypersensitive animals.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig1highres.jpg" width="500" /> 
Figure 1. A five-step procedure to monitor morphine induced-hyperalgesia and tolerance. The protocol is divided into five sequential steps (A-E) over a total duration of 15 days.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig2highres.jpg" width="500" /> 
Figure 2. Definition of the basal nociceptive response values (Step B; d-4 to d-1). The tail immersion (TIT) (A) and tail pressure (TPT) (B) tests are applied to the whole series of animals in order to evaluate their basal nociceptive values. Thereafter, two groups of mice (n=8), referred to as &#8216;Saline&#8217; and &#8216;Morphine&#8217; groups, are defined so that they exhibit stable and comparable average nociceptive values, whatever the nociceptive modality that is considered.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig3highres.jpg" width="500" /> 
Figure 3. Time-course for morphine analgesia at day 0 (Step C) in TIT (A) and TPT (B). The basal nociceptive response value of mice was determined every 30 min after single morphine (5 mg/kg, sc.) or saline injections. Data are expressed as means &#177; SEM, n=8 mice per group.*P &#60; 0.05, **P &#60; 0.01, ***P &#60; 0.001, unpaired t-test as compared to control group.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig4highres.jpg" width="500" /> 
Figure 4. Development of hyperalgesia following repeated morphine administration (Step D; d1 to d6). The basal nociceptive value of mice was determined by TIT (A) and TPT (B) once daily before morphine (5 mg/kg, sc.) or saline administration. Data are expressed as mean values &#177; SEM, n=8 mice per group. *P &#60; 0.05, **P &#60; 0.01, ***P &#60; 0.001 by unpaired t-test as compared with the saline-treated control group.
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig5highres.jpg" width="500" /> 
Figure 5. Time-course for morphine analgesia in chronic-morphine treated mice at day 7 (Step E) in TIT (A) and TPT (B). Mice that were chronically treated with morphine (black dots) or saline (white triangles) from day 0 to day 6, received on day 7 a single injection of morphine (5 mg/kg, sc.) or saline, respectively. The nociceptive response of mice was determined every 30 min after morphine or saline injection. Data are expressed as means &#177; SEM, n=8 mice per group. *P &#60; 0.05, **P &#60; 0.01 by unpaired t-test as compared with the saline-treated control group. Error bars that do not exceed symbols size are hidden.
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51264/51264fig6highres.jpg" width="500" /> 
Figure 6. Comparison of maximum analgesic responses of mice to morphine (5 mg/kg, sc.) before (day 0) and after chronic morphine treatment (day 7). Values reported here are from experiments shown in Figure 3 and Figure 5. Nociceptive values were measured using TIT (A) and TPT (B) 30 min after morphine or saline injection. Data are expressed as means &#177; SEM, n=8 mice per group. ***P &#60; 0.001 by paired t-test.

Watch this Scientific Journal Video about Assessment of Morphine-induced Hyperalgesia and Analgesic Tolerance in Mice Using Thermal and Mechanical Nociceptive Modalities at JoVE.com

Assessment of Morphine-induced Hyperalgesia and Analgesic Tolerance in Mice Using Thermal and Mechanical Nociceptive Modalities

We describe a protocol to examine the development of opioid-induced hyperalgesia and tolerance in mice. Based on the measurement of thermal and mechanical nociceptive responses of na&#239;ve and morphine-treated animals, it allows to quantify the increase in pain sensitivity (hyperalgesia) and decrease in analgesia (tolerance) associated with chronic opiate administration.

Opioid-induced hyperalgesia and tolerance severely impact the clinical efficacy of opiates as pain relievers in animals and humans. The molecular ...

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Research

JoVE Journal

Neuroscience

33.2K ビュー.  Université de Strasbourg. 私たちは、オピオイド誘発性痛覚過敏およびマウスにおける耐性の発生を調べるためのプロトコルを記述します。ナイーブおよびモルヒネで処置した動物の熱的および機械的な侵害刺激反応の測定に基づいて、疼痛感受性（痛覚過敏）の増加を定量化および慢性アヘン剤投与に関連した痛覚脱失（許容範囲）が減少することを可能にする。 

ビデオ: Assessment of Morphine-induced Hyperalgesia and Analgesic Tolerance in Mice Using Thermal and Mechanical Nociceptive Modalities

Assessment of Motor Balance and Coordination in Mice using the Balance Beam

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice. Fine motor coordination and balance can be assessed by the beam walking assay. The goal of this test is for the mouse to stay upright and walk across an elevated narrow beam to a safe platform. This test takes place over 3 consecutive days: 2 days of training and 1 day of testing. Performance on the beam is quantified by measuring the time it takes for the mouse to traverse the beam and the number of paw slips that occur in the process. Here we report the protocol used in our laboratory, and representative results from a cohort of C57BL/6 mice. This task is particularly useful for detecting subtle deficits in motor skills and balance that may not be detected by other motor tests, such as the Rotarod.

Deficits in fine motor coordination can be assessed with the balance beam test. Performance on the beam is quantified by the speed at which the beam is traversed and the number of times the mouse slips on the beam.

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice.  Fine motor coordination and ...

Rapid Determination of the Thermal Nociceptive Threshold in Diabetic Rats

Painful diabetic neuropathy (PDN) is characterized by hyperalgesia i.e., increased sensitivity to noxious stimulus, and allodynia i.e., hypersensitivity to normally innocuous stimuli1. Hyperalgesia and allodynia have been studied in many different rodent models of diabetes mellitus2. However, as stated by B&ouml;lcskei et al, determination of "pain" in animal models is challenging due to its subjective nature3. Moreover, the traditional methods used to determine behavioral responses to noxious thermal stimuli usually lack reproducibility and pharmacological sensitivity3. For instance, by using the hot-plate method of Ankier4, flinch, withdrawal and/or licking of either hind- and/or fore-paws is quantified as reflex latencies at constant high thermal stimuli (52-55 &deg;C). However, animals that are hyperalgesic to thermal stimulus do not reproducibly show differences in reflex latencies using those supra-threshold temperatures3,5. As the recently described method of B&ouml;lcskei et al.6, the procedures described here allows for the rapid, sensitive and reproducible determination of thermal nociceptive thresholds (TNTs) in mice and rats. The method uses slowly increasing thermal stimulus applied mostly to the skin of mouse/rat plantar surface. The method is particularly sensitive to study anti-nociception during hyperalgesic states such as PDN. The procedures described bellow are based on the ones published in detail by Alm&aacute;si et al 5 and B&ouml;lcskei et al 3. The procedures described here have been approved the Laboratory Animal Care and Use Committee (LACUC), Wright State University.

Here, we describe a rapid reliable and simple procedure to determine the lowest temperature at which rats or mice show nocifensive behavior, i.e. the thermal nociceptive threshold (TNT). This method applies a slowly increasing thermal stimulus allowing precise and reproducible estimation of TNTs with minimum, if any, stress to the animals.

Painful diabetic neuropathy (PDN) is characterized by hyperalgesia i.e., increased sensitivity to noxious stimulus, and allodynia i.e., hypersensitivity ...

Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity and extent of dendritic arborization correlates well with the synaptic plasticity in these cells. A reliable method for assessment of dendritic arborization is needed to characterize the deleterious effects of neurological disorders on these structures and to determine the effects of therapeutic interventions. However, quantification of these structures has proven to be a formidable task given their complex and dynamic nature. Fortunately, sophisticated imaging techniques can be paired with conventional staining methods to assess the state of dendritic arborization, providing a more reliable and expeditious means of assessment. Below is an example of how these imaging techniques were paired with staining methods to characterize the dendritic arborization in wild type mice. These complementary imaging methods can be used to qualitatively and quantitatively assess dendritic arborization that span a rather wide area within the hippocampal region.

We describe two methods for visualization and quantification of dendritic arborization in the hippocampus of mouse models: real-time and extended depth of field imaging. While the former method allows sophisticated topographical tracing and quantification of the extent of branching, the latter allows speedy visualization of the dendritic tree.

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation. There are several approaches to Golgi staining, all of which have been useful for determining the morphological characteristics of dendritic arbors and produce a clear background. The present Golgi-Cox method, (a slight variation of the protocol that is provided with a commercial Golgi staining kit), was designed to assess how a relatively low dose of the chemotherapeutic drug 5-flurouracil (5-Fu) would affect dendritic morphology, the number of spines, and the complexity of arborization within the hippocampus. The 5-Fu significantly modulated the dendritic complexity and decreased the spine density throughout the hippocampus in a region-specific manner. The data presented show that the Golgi staining method effectively stained the mature neurons in the CA1, the CA3, and the dentate gyrus (DG) of the hippocampus. This protocol reports the details for each step so that other researchers can reliably stain tissue throughout the brain with high quality results and minimal troubleshooting.

Here we present a Golgi-Cox protocol in extensive detail. This reliable tissue stain method allows for a high-quality assessment of the cytoarchitecture in the hippocampus, and throughout the entire brain, with minimal troubleshooting.

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching ...

Assessment of Dopaminergic Homeostasis in Mice by Use of High-performance Liquid Chromatography Analysis and Synaptosomal Dopamine Uptake

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) neurotransmission is strongly associated with several central nervous system-associated diseases such as Parkinson&#39;s disease, attention-deficit-hyperactivity disorder and drug addiction1,2,3,4. Delineating disease mechanisms involving DA imbalance is critically dependent on animal models to mimic aspects of the diseases, and thus protocols that assess specific parts of the DA homeostasis are important to provide novel insights and possible therapeutic targets for these diseases.
Here, we present two useful experimental protocols that when combined provide a functional read-out of the DAergic system in mice. Biochemical and functional parameters on DA homeostasis are obtained through assessment of DA levels and dopamine transporter (DAT) functionality5. When investigating the DA system, the ability to reliably measure endogenous levels of DA from adult brain is essential. Therefore, we present how to perform high-performance liquid chromatography (HPLC) on brain tissue from mice to determine levels of DA. We perform the experiment on tissue from dorsal striatum (dStr) and nucleus accumbens (NAc), but the method is also suitable for other DA-innervated brain areas.
DAT is essential for reuptake of DA into the presynaptic terminal, thereby controlling the temporal and spatial activity of released DA. Knowing the levels and functionality of DAT in the striatum is of major importance when assessing DA homeostasis. Here, we provide a protocol that allows to simultaneously deduce information on surface levels and function using a synaptosomal6 DA uptake assay.
Current methods combined with standard immunoblotting protocols provide the researcher with relevant tools to characterize the DAergic system.

Synaptosomal dopamine uptake and high-performance liquid chromatography analysis represent experimental tools to investigate dopamine homeostasis in mice by assessing the function of the dopamine transporter and levels of dopamine in striatal tissue, respectively. Here we present protocols to measure dopamine tissue content and assess the functionality of the dopamine transporter.

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) ...

Visual Evoked Potential Recordings in Mice Using a Dry Non-invasive Multi-channel Scalp EEG Sensor

For scalp EEG research environments with laboratory mice, we designed a dry-type 16 channel EEG sensor which is non-invasive, deformable, and re-usable because of the plunger-spring-barrel structural facet and mechanical strengths resulting from metal materials. The whole process for acquiring the VEP responses in vivo from a mouse consists of four steps: (1) sensor assembly, (2) animal preparation, (3) VEP measurement, and (4) signal processing. This paper presents representative measurements of VEP responses from multiple mice with a submicro-voltage signal resolution and sub-hundred millisecond temporal resolution. Although the proposed method is safer and more convenient compared to other previously reported animal EEG acquiring methods, there are remaining issues including how to enhance the signal-to-noise ratio and how to apply this technique with freely moving animals. The proposed method utilizes easily available resources and shows a repetitive VEP response with a satisfactory signal quality. Therefore, this method could be utilized for longitudinal experimental studies and reliable translational research exploiting non-invasive paradigms.

We designed a dry-type 16 channel EEG sensor which is non-invasive, deformable, and re-usable. This paper describes the whole process from manufacturing the proposed EEG electrode to signal processing of visual evoked potential (VEP) signals measured on a mouse scalp using a dry non-invasive multi-channel EEG sensor.

For scalp EEG research environments with laboratory mice, we designed a dry-type 16 channel EEG sensor which is non-invasive, deformable, and re-usable ...

Assessment of Sensory Thresholds in Dogs Using Mechanical and Hot Thermal Quantitative Sensory Testing

Quantitative sensory testing (QST) is used to evaluate the function of the somatosensory system in dogs by assessing the response to applied mechanical and thermal stimuli. QST is used to determine normal dogs&#39; sensory thresholds and evaluate alterations in peripheral and central sensory pathways caused by various disease states, including osteoarthritis, spinal cord injury, and cranial cruciate ligament rupture. Mechanical sensory thresholds are measured by electronic von Frey anesthesiometers and pressure algometers. They are determined as the force at which the dog exhibits a response indicating conscious stimulus perception. Hot thermal sensory thresholds are the latency to respond to a fixed or ramped temperature stimulus applied by a contact thermode.
Following a consistent protocol for performing QST and paying attention to details of the testing environment, procedure, and individual study subjects are critical for obtaining accurate QST results for dogs. Protocols for the standardized collection of QST data in dogs have not been described in detail. QST should be performed in a quiet, distraction-free environment that is comfortable for the dog, the QST operator, and the handler. Ensuring that the dog is calm, relaxed, and properly positioned for each measurement helps produce reliable, consistent responses to the stimuli and makes the testing process more manageable. The QST operator and handler should be familiar and comfortable with handling dogs and interpreting dogs&#39; behavioral responses to potentially painful stimuli to determine the endpoint of testing, reduce stress, and maintain safety during the testing process.

This work describes a standard protocol for mechanical and hot thermal quantitative sensory testing to evaluate the somatosensory system in dogs. Sensory thresholds are measured using an electronic von Frey anesthesiometer, pressure algometer, and hot contact thermode.

Quantitative sensory testing (QST) is used to evaluate the function of the somatosensory system in dogs by assessing the response to applied mechanical ...

Recording Network Activity in Spinal Nociceptive Circuits Using Microelectrode Arrays

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an increasingly detailed view of the circuits underpinning spinal pain processing. However, the effects of these connections for broader network activity in the DH remain less well understood because most studies focus on the activity of single neurons and small microcircuits. Alternatively, the use of microelectrode arrays (MEAs), which can monitor electrical activity across many cells, provides high spatial and temporal resolution of neural activity. Here, the use of MEAs with mouse spinal cord slices to study DH activity induced by chemically stimulating DH circuits with 4-aminopyridine (4-AP) is described. The resulting rhythmic activity is restricted to the superficial DH, stable over time, blocked by tetrodotoxin, and can be investigated in different slice orientations. Together, this preparation provides a platform to investigate DH circuit activity in tissue from na&#239;ve animals, animal models of chronic pain, and mice with genetically altered nociceptive function. Furthermore, MEA recordings in 4-AP-stimulated spinal cord slices can be used as a rapid screening tool to assess the capacity of novel antinociceptive compounds to disrupt activity in the spinal cord DH.

The combined use of microelectrode array technology and 4-aminopyridine-induced chemical stimulation for investigating network-level nociceptive activity in the spinal cord dorsal horn is outlined.

The roles and connectivity of specific types of neurons within the spinal cord dorsal horn (DH) are being delineated at a rapid rate to provide an ...

Mechanical Conflict-Avoidance Assay to Measure Pain Behavior in Mice

Pain comprises of both sensory (nociceptive) and affective (unpleasant) dimensions. In preclinical models, pain has traditionally been assessed using reflexive tests that allow inferences regarding pain's nociceptive component but provide little information about the affective or motivational component of pain. Developing tests that capture these components of pain are therefore translationally important. Hence, researchers need to use non-reflexive behavioral assays to study pain perception at that level. Mechanical conflict-avoidance (MCA) is an established voluntary non-reflexive behavior assay, for studying motivational responses to a noxious mechanical stimulus in a 3 chamber paradigm. A change in a mouse's location preference, when faced with competing noxious stimuli, is used to infer the perceived unpleasantness of bright light versus tactile stimulation of the paws. This protocol outlines a modified version of the MCA assay which pain researchers can use to understand affective-motivational responses in a variety of mouse pain models. Though not specifically described here, our example MCA data use the intraplantar complete Freund's adjuvant (CFA), spared nerve injury (SNI), and a fracture/casting model as pain models to illustrate the MCA procedure.

The mechanical conflict-avoidance assay is used as a non-reflexive readout of pain sensitivity in mice which can be used to better understand affective-motivational responses in a variety of mouse pain models.

Pain comprises of both sensory (nociceptive) and affective (unpleasant) dimensions. In preclinical models, pain has traditionally been assessed using ...