Name: a method to study the impact of chemically-induced ovarian failure on exercise capacity and cardiac adaptation in mice
Uploaded: 2014-04-07T15:00:00
Duration: 14 min 26 s
Description: Watch this Scientific Journal Video about A Method to Study the Impact of Chemically-induced Ovarian Failure on Exercise Capacity and Cardiac Adaptation in Mice at JoVE.com

This work was supported by NIH grant (HL 098256), by a National and Mentored Research Science Development Award (K01 AR052840) and Independent Scientist Award (K02 HL105799) from the NIH awarded to J.P. Konhilas and the Interdisciplinary Training Grant in Cardiovascular Sciences (HL007249). Support was received from the Sarver Heart Center at the University of Arizona and from the Steven M. Gootter Foundation.

All experiments, housing, and caring for animals was performed using protocols that adhered to guidelines and approved by the Institutional Animal Care and Use Committee at the University of Arizona, and to 2011 NIH guidelines for care and use of laboratory animals.&#160;
1. Treatment with 4-Vinylcyclohexene Diepoxide
<ol>
	<li>VCD dosing protocol: In a chemical fume hood, add 0.587 ml of VCD to a clean flask, bring the final volume to 10 ml with sesame oil, and then mix by inverting gently, cover&#160;and seal with Parafilm. Transfer the solution to a dosing vial, seal it with Parafilm, and store at 4 &#176;C for up to 7 days. Be sure all reagents are sterile.&#160;
	<ol>
		<li>Use sesame oil as the vehicle control as VCD is dissolved in sesame oil. Store VCD (MW=140.2 g/mol, density=1.09 g/ml) at -20 &#176;C.</li>
		<li>VCD injection protocol: At 2 months of age, weigh and administer daily intraperitoneal injections of room temperature VCD at a dose of 160 mg/kg for 20 consecutive days. Inject control mice in a similar manner with the vehicle solution, sesame oil, at a volume of 2.5 ml/kg. 
		Note: During the injection period, try to alternate the sites of injection to reduce the chances of local infection. Use the following equation to calculate the volume needed for injection: Body weight (g) x kg/1,000 g x 2.5 ml/kg = volume (ml) of solution to inject resulting in approximately 50 ml/injection using a 27 G needle. Due to the size of the needle local anesthetics are not needed.</li>
		<li>Estrous cycle monitoring: After the injection period is complete, monitor the estrous cycles daily by obtaining vaginal smears. Restrain mice using the dorsum scruff method. Insert an eyedropper with 0.2 ml of sterile saline into the vaginal orifice to a depth of 0.5 cm. Triturate saline (3x) and place 1 drop onto a blank slide. Immediately evaluate samples microscopically using a standard light microscope at 100X&#160;magnitude. 
		Note: Vaginal cytology is determined to ensure loss of cycling and therefore validate ovarian failure. Figure 2A illustrates typical vaginal cytology across different phases of the estrous cycle in mice, namely proestrus, estrus, metestrus, diestrus17-19. During proestrus, cells include nucleated epithelial cells (Figure 2A, arrows indicate nucleated cells); during estrus, cells are predominantly cornified squamous epithelial cells; Metestrus is characterized by a mix of cell types, with a predominance of leukocytes; Diestrus is dominated by leukocytes. Adjustment of the focal plane may be required to visualize cells and nuclei throughout the sample. As illustrated in Figure 2B, a typical murine estrus cycle is 4-5 days. Mice are considered acyclic after 15 consecutive days in persistent diestrus. Continuously monitor estrous cycles throughout the duration of the study period to confirm lack of estrous cycles in VCD treated mice.</li>
	</ol>
	</li>
</ol>
2. Voluntary Cage-wheel Exercise
 
Inbred C57BL/6 and B6C3F1 female mice, aged 7 months, are exposed to voluntary cage wheel running. House animals individually in a cage (47 cm x&#160;26 cm x&#160;14.5&#160;cm) with unencumbered access to a cage wheel for 28 days.
<ol>
	<li>Equipment setup: The cage wheel apparatus consists of an 11.5 cm&#160;diameter wheel with a 5.0 cm&#160;wide running surface equipped with a digital magnetic counter that is activated by wheel rotation. Before each wheel running test, calibrate the cage wheel according to the manufacturer&#39;s guidelines. Measure the inner diameter of the running wheel, calculate the circumference and enter this value into the digital counter as wheel size.
	<ol>
		<li>Attach the computer portion of the digital magnetic counter to the wheel stand using tape.</li>
		<li>Attach the computer portion of the digital magnetic counter to the wheel stand using tape.</li>
		<li>Attach the free magnet of the digital magnetic counter to the central bar on the same side as the magnet cabled to the computer, again taping sufficiently to protect the magnet from chewing damage. The magnets should be positioned so that each rotation of the wheel is recorded by the computer without any interference or resistance to complete wheel rotation. 
		Note: Duct tape has been used successfully to attach the components of the digital magnetic counter to the running wheel. Taping should be sufficient to completely cover the magnet and prevent damage from chewing by the animal. The magnet should be positioned as close as possible to the wheel without hindering its movement. However, the setup should be monitored daily for chewing damage and repaired as necessary.</li>
		<li>Mount the wheel to the cage by sliding the wheel stand through the top wiring of the animal cage. Secure the wheel stand to the cage wiring with binder clips. 
		Note: The clip should be on the top of the wiring, away from the animal. Feed the computer portion and any extra cable through the wiring of the cage. Place the computer and excess cable out of reach of the animal. This may be accomplished by placing a medium-sized plastic weigh dish on top of the wiring of the cage to hold the computer and excess cable.</li>
	</ol>
	</li>
	<li>Exercise: For a given litter, randomly assign mice to either the sedentary control or exercise regimen. Give all animals water and standard hard rodent chow ad libitum.
	<ol>
		<li>Manually record daily values for the distance ran (km/day) and total time spent running (hr/day), given by the magnetic counter. Record distance and time values within the same 1 hr time frame every day for accurate and consistent values throughout the 28 day duration of the exercise period. Clear (zero) the magnetic counter after each day&#39;s recording to ensure accurate readings the following day.</li>
		<li>Calculate the average values of distance and speed for each group. Calculate speed based on the given values of distance and time each day for each mouse. Data can be displayed as daily or weekly averages for each measured parameter.</li>
	</ol>
	</li>
	<li>Tissue harvest: At the end of the specific exercise period, euthanize exercised and sedentary control animals within 30 min of removal from cages by cervical dislocation under inhaled anesthesia. Weigh body mass and rapidly excise hearts, skeletal muscle, and ovaries.
	<ol>
		<li>At the time of tissue harvesting, mount and stain the collected ovaries with hemotoxylin and eosin (H&#38;E; Figure 2C). Under magnification, count and classify follicles as primordial, primary, secondary, or antral for each VCD-treated and control animal20. A primordial follicle contains a small primary oocyte, a single layer of flattened or squamous granulosa cells closely opposed to the oocyte, and a basal lamina. A primary follicle is defined by the presence of one or more cuboidal granulosa cells that are arranged in a single layer surrounding the oocyte. A secondary follicle contains granulosa cells that form multiple layers around the oocyte, and a theca. An antral follicle is characterized by a cavity or &#8220;antrum&#8221; containing fluid.</li>
		<li>To excise the heart, make a medial incision on the ventral side and visualize the heart after opening the diaphragm. Rapidly excise the heart at the base and place in cold phosphate buffered saline to sufficiently wash-out residual ventricular blood. Trim the heart of excess tissue, weigh the whole heart and snap-freeze in liquid nitrogen.</li>
		<li>Wash all other excised tissues with a modified ice-cold phosphate-buffered saline solution (PBS), and snap-freeze in isopentane cooled in liquid nitrogen.</li>
	</ol>
	</li>
</ol>
3. Forced Treadmill Exercise
<ol>
	<li>When employing a forced exercise paradigm to measure exercise capacity, use a treadmill and set up as follows:
	<ol>
		<li>Place an electrical shock grid at the base of the treadmill ramp. This shock generated serves as a stimulus for exercise. The intensity and frequency of the electrical stimuli are controlled manually by the users. Grids can be enabled or disabled individually for each lane. All data collection is done manually.</li>
		<li>Periodically, calibrate the treadmill according to the manufacturer&#39;s guidelines.
		<ol>
			<li>Ensure that the Treadmill Controller front panel toggle is set to &#8220;STOP&#8221;.</li>
			<li>Place speed sensor on treadmill belt lane1 and securely fasten with screw nut in position shown below.</li>
			<li>Using the dial on the Treadmill Controller front panel, adjust the speed to 88.7 m/min Then, set the electric stimulus frequency to 2 Hz on Treadmill Controller front panel.</li>
			<li>Push the &#8220;ODOMETER RESET&#8221; button to initialize calibration procedure. 
			Note: Sedentary or nonexercise control groups must be included in the experimental design. The sedentary group is age matched to the exercise group. We have previously shown that housing the sedentary mice individually in cages of equal size to the exercised mice did not impact the parameters of interest when compared to sedentary mice housed with siblings. Therefore, sedentary mice were kept with siblings in the same room as the exercised mice.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Acclimation: Acclimate mice to the treadmill 3-7 days prior to the experiment daily. To do this, place mice in the treadmill with the belt unmoving and shock grids off, but with the belt motor on (noise). Leave mice undisturbed for at least 15 min.</li>
	<li>Warm up: Warm up steps as used in this protocol are as follows:
	<ol>
		<li>Place each mouse on the treadmill. Disable the shock grid and increase belt speed to 4 m/min for 15 min.</li>
		<li>Enable the shock grid and maintain belt speed at 4 m/min belt speed for an additional 10 min.</li>
	</ol>
	</li>
	<li>Exercise training regimens
	<ol>
		<li>Following warm up, keep shock grids on and increase belt speed to 5 m/min for 15 min.</li>
		<li>Increase belt speed to 15 m/min for an additional 15 min.</li>
		<li>Following training, transfer mice back to their cages. 
		Note: In this study, exercise training was performed daily for 1 week.</li>
		<li>Acute exercise regimen: An acute exercise regimen is designed to determine the short and long term effects of extreme exercise stress.
		<ol>
			<li>Following warm up, increase belt speed from 5 m/min to 10, 15, 20, 25, and 30 m/min in 10 min intervals. 
			Note: Terminate exercise if mice are unable to continue running or show signs of exhaustion or distress. In our study, most animals were stopped at 25 m/min, and no animal passed the 30 m/min stage.</li>
			<li>Test each mouse three times, allowing 2-3 days between each test. Calculate average values across all exercise sessions for each mouse.</li>
		</ol>
		</li>
		<li>Exercise regimen for endurance: An endurance exercise protocol is designed to train mice for an extended exposure to the treadmill exercise stimulus. This type of protocol tests the capacity to tolerate the long-term effects of moderate exercise stress.
		<ol>
			<li>Begin with the belt speed outlined during the warm up period, 4 m/min.</li>
			<li>Gradually increase belt speed by 1m/min until a final speed of 20 m/min is reached. Total duration of a typical study protocol is 50 min.</li>
			<li>After exercise regimen, transfer mice back to their cages. 
			Note: Running mice to exhaustion is avoided. If at any point during the experiment a mouse shows signs of exhaustion, deactivate the shock grid and allow the mouse to rest and recover.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Data and Statistical Analysis
<ol>
	<li>Present results as mean &#177; SEM and use 2-way ANOVA (treatment group and exercise as main factors) followed by a Student-Newman Keuls post hoc test or a student&#39;s t test to compare differences between mean values.</li>
	<li>Determine the percent change in cardiac mass with exercise by comparing the mass of each exercised animal to the mean cardiac mass of the sedentary group. The difference in cardiac mass is then expressed as a percent change from sedentary animals for each respective animal. p values of &#60;0.05 are considered statistically significant.</li>
</ol>

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The authors have nothing to disclose.

Both cage wheel exercise and treadmill running exercise regimens have been used in our previous studies16,21,22. All animals are kept in dedicated exercise rooms to minimize environmental or human disturbances. Any disturbance of the animals&#39; biological rhythm can significantly affect animals&#39; exercise performance. Therefore, the entire animal care, performance monitoring and recording, and exercise regimens should be done within the same daily time frame. Investigators should strive to avoid extra stress to the animals by exhibiting precautions such as limiting the duration of stress exposure and covering the cages when appropriate.
Cage wheel exercise
When preparing the cage wheel apparatus, the wheel should be easily accessible for the mouse to mount and exit. There should minimal bedding at the bottom of the cage to prevent mice from nesting around the wheel and hindering free wheel movement. The magnetic sensor was placed on the stationary portion of the wheel and the magnet secured to the free wheel, so as to count each rotation of the wheel as the mouse exercised. The recording device attached to the magnetic sensor should be placed above the wire lid of the cage, and all exposed wires need to be either taped or kept out of reach from the mice, so as to provide a barrier against chewing damage. The magnet and magnetic sensor were also taped to the appropriate structures to prevent both damage and unwanted movement. During daily data collection, all the taped wires must be closely checked to insure the wires remain intact and should be retaped as necessary. Since it may require several days for mice to acclimatize to the new environment and begin exercising regularly, data recording should be tailored to each individual animal. Although animals tend to spend many hr on the wheel, most of the running occurs at night. Therefore, some animals may reveal high performance activity even if they appear dormant during daytime. Furthermore, it is not uncommon that some animals will exhibit limited cage wheel running. Typically, cage wheel activity that amounts to an average 1 km/day or less is the result of random cage activity and not associated with any significant running stimulus. Therefore, animals whose running parameters are at 1 km/day or 1 hr/day or less should be excluded in the final data analysis nonrunners.
Depending on the specific purposes of an individual study and strains of mice involved, the length of exercise can be modified. Our previous study showed that for both mouse strains involved, the average wheel-running speeds and the duration of exercise on the cage wheel gradually increased over time and plateaued by 4 weeks.
Cage wheel running is a type of voluntary exercise in animal models, which is generally taken as being performed under less stressful conditions. Unlike treadmill exercise in which certain parameters can be preset, all running in the test cage is voluntary and easily affected by environmental or behavioral cues. Therefore, the comparisons between different experimental settings are typically avoided unless large experimental groups numbers are used. Other potential areas of research that can benefit from this type of exercise paradigm are studies that incorporate behavioral responses with physiological consequences such as cognitive neuroscience. For studies utilizing the voluntary cage wheel paradigm, the most critical steps are cage set up and monitoring of the exercise activities.
Treadmill exercise 
Acclimation steps are necessary to prepare animals for the treadmill exercise protocol. Depending on the specific purpose of the individual study, the duration of the acclimation period can vary. In this study, a period of one week was chosen to allow animals to acclimate to the treadmill apparatus and motor noise in the cage, warm&#160;up periods are an important step of the exercise training regimen. Similar to human counterparts, the mice must be warmed up before getting into any intensive exercise regimens to avoid injury and potential data artifacts. In this study, we began the belt at the lowest speed possible (4 m/min) with the shock grids turned off for 15 min. All mice involved started to run on and off the belt. The shock grids were then turned on and the belt speed maintained at 4 m/min. To avoid over-stressing the mice, we set the electric shock at a mild level (1 Hz). Within a few days, all mice adapted to the treadmill belt.
The duration of running as well as the acceleration can also be tailored to each experimental design. A recommended starting acceleration rate is 1 m/min. The treadmill lanes of mice showing signs of exhaustion must be turned off immediately. Typically signs of exhaustion exhibited by mice on the treadmill are as follows:
<ol>
	<li>Continuous exposure to the shock grid for &#62;5-10 sec,</li>
	<li>Loss of running capacity resulting in shock grid exposure &#62;5x during a single exercise regimen, and</li>
	<li>Loss of running capacity resulting in shock grid exposure &#62;2 sec at least 3x during a single exercise regimen.</li>
</ol>
The values listed above are to be used primarily as guidelines for a specific experiment. These parameters need to be determined for every exercise protocol as it varies from mouse strain to mouse strain, experiment to experiment, and investigator to investigator.
Unlike cage wheel exercise, treadmill studies must be repeated several times to eliminate the effects of environmental and biological factors on running performance. It is not uncommon to see fluctuations in running performance between tests even for the same animal. Since each exercise regimen exposes running mice to a stress, an appropriate resting interval between tests is necessary. In our experience, 2-3 days resting period between each test was sufficient for mice to recover.
Like human treadmill exercise, the speed for endurance exercise is best determined by the heart rate or blood oxygen saturation, a technically difficult feat with mouse studies. The most critical step of treadmill studies is determination of exhaustion, which usually varies between individual mice, and was set as 80% maximal speed. In our own experience the recording of results may significantly differ between individual researchers, since the criteria for exhaustion are mostly subjective. Thus, the consistent application of the same criteria during the recording is critical.

The natural onset of menopause is characterized by the depletion of ovarian primordial follicles through atresia, eventually resulting in ovarian senescence. In the United States, over 30% of a women&#39;s lifetime will be spent post-menopausal. In response to a decrease in ovarian function, several physiological and psychological consequences may occur, including increased adiposity, vascular instability, and mood or sleep disturbances. Because of increased obesity/adiposity compared with premenopausal counterparts1,2, postmenopausal women are particularly more susceptible to metabolic syndrome and associated comorbidities, including cardiovascular disease (CVD)3.&#160;Moreover, it is becoming increasingly evident that successful cardiac rehabilitation for patients with CVD includes regular aerobic exercise, and that exercise reduces cardiovascular morbidity and mortality in these subjects4,5.&#160;Yet, how exercise capacity changes during the pre- to post-menopausal transition has not been well studied. More importantly, the role of exercise as a preventative measure for CVD risk in post-menopausal women remains remarkably understudied.
The most commonly used inducible model for mimicking menopause in women is the ovariectomized (OVX) rodent. Recently, the occupational chemical 4-vinylcyclohexene diepoxide (VCD) has been shown to specifically target small primary and primordial ovarian follicles by accelerating the natural process of atresia, ultimately resulting in ovarian failure with no obvious toxicity found in other tissues6. The estrous cycle of VCD-treated mice, which is analogous to a human menstrual cycle, ceases within 2-3 months after the completion of VCD injections, which in turn results in a gradual withdrawal of estrogen, mimicking the peri- to post-menopausal transition. Thus, a follicle-deplete, ovary-intact animal closely approximates the natural human progression through the pre- to peri- to post-menopausal transition7-9.&#160;Moreover, it provides a nonsurgical alternative to the OVX model reducing complications from surgery, such as infection. In this study, we employed VCD-induced menopausal mice to study the impact of exercise on the cardiac adaption in menopausal mice.
There are several instances that suggest the association between estrogen and cage wheel exercise in rodents of both sexes10-13. Estrogen depletion by surgical OVX decreases voluntary exercise activity in mice and rats14,15. Two methods of exercise were used in this study to test the exercise capacity of VCD-induced menopausal mice. Cage wheel running is generally regarded as a type of voluntary exercise in animal models and performed, presumably, under less stressful conditions. However, cage wheel running is not indicative of the relative exercise capacity of animals, which requires animals to exercise at much higher levels. Forced treadmill exercise was used to determine exercise capacity, measured as a tolerance to exercise intensity and endurance. In addition, we have previously shown that cage wheel exercise provides a stimulus for cardiac hypertrophy and this hypertrophic growth is sex-dependent16.&#160;Therefore, we also measured cardiac adaption to cage wheel exercise in VCD-induced menopausal mice.

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	<colgroup>
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr>
			<td>4-vinylcyclohexene diepoxide (VCD)</td>
			<td>Sigma, cat# V-3630</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>11.5-cm-diameter wheel with a 5.0-cm-wide running surface</td>
			<td>Petsmart,model 6208</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Digital magnetic counter</td>
			<td>Sigma Sport,model BC 600</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Treadmill exercise</td>
			<td>Columbus Instruments , model 1055SDRM</td>
			<td></td>
			<td></td>
		</tr>
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a method to study the impact of chemically-induced ovarian failure on exercise capacity and cardiac adaptation in mice

The risk of cardiovascular disease (CVD) increases in post-menopausal women, yet, the role of exercise, as a preventative measure for CVD risk in post-menopausal women has not been adequately studied. Accordingly, we investigated the impact of voluntary cage-wheel exercise and forced treadmill exercise on cardiac adaptation in menopausal mice. The most commonly used inducible model for mimicking menopause in women is the ovariectomized (OVX) rodent. However, the OVX model has a few dissimilarities from menopause in humans. In this study, we administered 4-vinylcyclohexene diepoxide (VCD) to female mice, which accelerates ovarian failure as an alternative menopause model to study the impact of exercise in menopausal mice. VCD selectively accelerates the loss of primary and primordial follicles resulting in an endocrine state that closely mimics the natural progression from pre- to peri- to post-menopause in humans. To determine the impact of exercise on exercise capacity and cardiac adaptation in VCD-treated female mice, two methods were used. First, we exposed a group of VCD-treated and untreated mice to a voluntary cage wheel. Second, we used forced treadmill exercise to determine exercise capacity in a separate group VCD-treated and untreated mice measured as a tolerance to exercise intensity and endurance.

A typical experimental protocol, such as that used in this study, is illustrated in Figure 1. Following 20 consecutive days of VCD treatment, vaginal cytology was used to determine the presence or absence of cyclicity in the VCD-injected mice; vehicle-injected mice were also monitored. The stage in the estrus cycle was determined by the proportion of epithelial cells, cornified epithelial cells, and leukocytes in the vaginal smear (Figure 2A). The presence of only cornified epithelial cells indicated the mouse was in estrus, and therefore was still cycling. The estrous cycle in mice typically lasts 4 days, therefore VCD injected animals were considered acyclic after 15 days of persistent diestrus, in which vaginal cytology revealed a majority of leukocytes (Figure 2A). The estrous cycle of VCD-treated mice became irregular, and then ceased within 2-3 months after the start of VCD injections (Figure 2B). VCD ovarian tissue showed significant atrophy, compared to controls. At high magnification, we see a complete depletion of primary and primordial follicles (Figure 2C).
To test the effect of VCD-induced ovarian failure on exercise performance, we randomized 7 month old VCD- and vehicle-treated mice (two strains, C57BL/6 and B6C3F1) to either sedentary or cage wheel exercise groups 4-6 weeks following the cessation of cycling (Figure 1). Daily exercise values were recorded for time and distance for each exercised animal throughout the 4&#160;week exercise period. Weekly averages of daily running values for distance, time and speed are shown in Figure 3 for C57BL/6 mice treated with vehicle or VCD. We did not see significant differences in exercise performance measured by average daily time, distance, and speed on the cage wheel between VCD- and vehicle-treated groups in either C57BL/6 or B6C3F1 mouse strain (Figure 4A). The calculated wheel-running speeds gradually increased over the 4-week running period as previously shown16, but were not significantly different between experimental groups. These studies indicate the impact of VCD-induced ovarian failure on voluntary cage wheel exercise transcends at least 2 mouse strains.
Following the duration of the voluntary cage wheel exercise protocol, body morphometrics were recorded and the hearts were rapidly excised and weighed. We have previously shown that voluntary cage wheel exercise induces an increase in cardiac mass16. Here, we also show cardiac hypertrophy in response to cage wheel exercise measured by absolute heart weight (HW) and HW normalized to tibial length (TL) (Table 1). The absolute heart mass and HW/TL ratio were significantly greater in vehicle- and VCD-treated mice when compared to sedentary counterparts. However, there were no measurable differences in cardiac hypertrophy between control and VCD-induced ovarian failure mice following voluntary cage wheel exercise.
Next, we used involuntary (forced) treadmill running to determine exercise capacity. We first subjected mice to a high-intensity protocol that increased average speed stepwise every 10 min until exhaustion. On average, the maximal speed of vehicle treated mice was 26.7&#177; 0.95 m/min and the maximal speed of VCD treated mice was 28&#177;1.4 m/min before exhaustion. We used lower-intensity running at 20 m/min (about 80% of maximal) to assay for endurance capacity. As shown in Figure 4B, no significant difference in high or low intensity exercise capacity was found between vehicle and VCD treated group, indicating that VCD-induced ovarian failure had no effect on mouse exercise capacity.
In conclusion, menopause induced by VCD treatment has no effect on either voluntary or forced running capacity, as well as the cardiac adaptive response to exercise.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51083/51083fig1highres.jpg" src="/files/ftp_upload/51083/51083fig1.jpg" /> 
Figure 1. VCD-administration and experimental protocol.&#160;Mice were administered VCD (160 mg/kg) or sesame oil (vehicle control) starting at 2 months of age for 20 consecutive days. Following VCD or vehicle injections, estrous cycles were monitored and cyclicities were determined as described in the Methods section. Loss of ovarian function occurred within 60-90 days and was confirmed by vaginal cytology. At 7 months of age, mice were exposed to voluntary cage wheel exercise for 4 weeks or subjected to the treadmill exercise protocol. 
<img alt="Figure 2" fo:content-width="3.5in" fo:src="/files/ftp_upload/51083/51083fig2highres.jpg" src="/files/ftp_upload/51083/51083fig2.jpg" /> 
Figure 2. VCD-induced ovarian failure in mice.&#160;A: Cytology of an unstained vaginal smear from a vehicle injected mouse demonstrating the four stages of the estrous cycle: 1) proestrous, predominantly consisting of nucleated epithelial cells 2) Estrus, characterized by un-nucleated cornified cells; 3) metestrus, consisting of nucleated epithelial cells (arrows), un-nucleated cornified epithelial cells (arrows), and leukocytes; 4) diestrus, consisting primarily of leukocytes; and. B: Average length (days) of the estrous cycle of vehicle- and VCD-injected groups. One cycle is measured from the first day in estrus to the first day of the next period in estrus. Vaginal cytology began approximately 6 weeks following the onset of injections. The estrous cycles of VCD-injected animals become irregular, compared to vehicle-injected animals, and after 12 estrus cycles, cytology of all VCD injected animals no longer indicated cycling. C: H&#38;E staining of ovarian tissue at 8 months of age. Cross-sections done at 5&#956;m thickness for vehicle-treated (left panel) and VCD-treated (right panel) mice at 20X&#160;(top panel) and 200X&#160;(bottom panel) magnification. Ovarian tissue at 200X&#160;magnification shows a number of follicles in vehicle- compared to VCD-treated mice. <a href="https://www.jove.com/files/ftp_upload/51083/51083fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51083/51083fig3highres.jpg" src="/files/ftp_upload/51083/51083fig3.jpg" /> 
Figure 3. Voluntary cage wheel performance in vehicle- or VCD-treated C57Bl/6J female mice. Left Panel: Average running distance (km/day) for every 1&#160;week period over the 4&#160;week study period. Middle Panel: Average running time (hr/day) for every 1&#160;week period over the 4&#160;week study period. Right Panel: Average running speed (km/hr) for every 1&#160;week period over the 4&#160;week study period. <a href="https://www.jove.com/files/ftp_upload/51083/51083fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/51083/51083fig4highres.jpg" src="/files/ftp_upload/51083/51083fig4.jpg" /> 
Figure 4. Voluntary cage wheel and treadmill performance in vehicle- and VCD-treated mice. A: (Top Panel) C57BL/6 average running distance (km/day) and time spent on the wheel (hrs/day) for every 24 hr over the 4 week study period.&#160;(Bottom Panel)&#160;B6C3F1 average running distance (km/day) and time spent on the wheel (hr/day) for every 24 hr over the 4 week study period. B: C57BL/6 treadmill running parameters. Maximum time (Top Panel) and maximum speed (Botoom Panel) using protocols detailed in the methods section.

Watch this Scientific Journal Video about A Method to Study the Impact of Chemically-induced Ovarian Failure on Exercise Capacity and Cardiac Adaptation in Mice at JoVE.com

A Method to Study the Impact of Chemically-induced Ovarian Failure on Exercise Capacity and Cardiac Adaptation in Mice

Two exercise paradigms were tested on a newly developed chemically induced menopausal mouse model to examine the impact of menopause on exercise capacity and cardiac adaption to exercise.

The risk of cardiovascular disease (CVD) increases in post-menopausal women, yet, the role of exercise, as a preventative measure for CVD risk in ...

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Phosphorus-31 Magnetic Resonance Spectroscopy: A Tool for Measuring In Vivo Mitochondrial Oxidative Phosphorylation Capacity in Human Skeletal Muscle

Skeletal muscle mitochondrial oxidative phosphorylation (OXPHOS) capacity, which is critically important in health and disease, can be measured in vivo and noninvasively in humans via phosphorus-31 magnetic resonance spectroscopy (31PMRS). However, the approach has not been widely adopted in translational and clinical research, with variations in methodology and limited guidance from the literature. Increased optimization, standardization, and dissemination of methods for in vivo 31PMRS would facilitate the development of targeted therapies to improve OXPHOS capacity and could ultimately favorably impact cardiovascular health. 31PMRS produces a noninvasive, in vivo measure of OXPHOS capacity in human skeletal muscle, as opposed to alternative measures obtained from explanted and potentially altered mitochondria via muscle biopsy. It relies upon only modest additional instrumentation beyond what is already in place on magnetic resonance scanners available for clinical and translational research at most institutions. In this work, we outline a method to measure in vivo skeletal muscle OXPHOS. The technique is demonstrated using a 1.5 Tesla whole-body MR scanner equipped with the suitable hardware and software for 31PMRS, and we explain a simple and robust protocol for in-magnet resistive exercise to rapidly fatigue the quadriceps muscle. Reproducibility and feasibility are demonstrated in volunteers as well as subjects over a wide range of functional capacities.

Phosphorus-31 Magnetic Resonance Spectroscopy: A Tool for Measuring In Vivo Mitochondrial Oxidative Phosphorylation Capacity in Human Skeletal Muscle

This work demonstrates the feasibility of an in vivo phosphorus-31 magnetic resonance spectroscopy (31PMRS) technique to quantify mitochondrial oxidative phosphorylation (OXPHOS) capacity in human skeletal muscle.

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Impact of Intracardiac Neurons on Cardiac Electrophysiology and Arrhythmogenesis in an Ex Vivo Langendorff System

Since its invention in the late 19th century, the Langendorff ex vivo heart perfusion system continues to be a relevant tool for studying a broad spectrum of physiological, biochemical, morphological, and pharmacological parameters in centrally denervated hearts. Here, we describe a setup for the modulation of the intracardiac autonomic nervous system and the assessment of its influence on basic electrophysiology, arrhythmogenesis, and cyclic adenosine monophosphate (cAMP) dynamics. The intracardiac autonomic nervous system is modulated by the mechanical dissection of atrial fat pads-in which murine ganglia are located mainly&#8212;or by the usage of global as well as targeted pharmacological interventions. An octapolar electrophysiological catheter is introduced into the right atrium and the right ventricle, and epicardial-placed multi-electrode arrays (MEA) for high-resolution mapping are used to determine cardiac electrophysiology and arrhythmogenesis. F&#246;rster resonance energy transfer (FRET) imaging is performed for the real-time monitoring of cAMP levels in different cardiac regions. Neuromorphology is studied by means of antibody-based staining of whole hearts using neuronal markers to guide the identification and modulation of specific targets of the intracardiac autonomic nervous system in the performed studies. The ex vivo Langendorff setup allows for a high number of reproducible experiments in a short time. Nevertheless, the partly open nature of the setup (e.g., during MEA measurements) makes constant temperature control difficult and should be kept to a minimum. This described method makes it possible to analyze and modulate the intracardiac autonomic nervous system in decentralized hearts.

Impact of Intracardiac Neurons on Cardiac Electrophysiology and Arrhythmogenesis in an Ex Vivo Langendorff System

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The Generation of Closed Femoral Fractures in Mice: A Model to Study Bone Healing

Bone fractures impose a tremendous socio-economic burden on patients, in addition to significantly affecting their quality of life. Therapeutic strategies that promote efficient bone healing are non-existent and in high demand. Effective and reproducible animal models of fractures healing are needed to understand the complex biological processes associated with bone regeneration. Many animal models of fracture healing have been generated over the years; however, murine fracture models have recently emerged as powerful tools to study bone healing. A variety of open and closed models have been developed, but the closed femoral fracture model stands out as a simple method for generating rapid and reproducible results in a physiologically relevant manner. The goal of this surgical protocol is to generate unilateral closed femoral fractures in mice and facilitate a post-fracture stabilization of the femur by inserting an intramedullary steel rod. Although devices such as a nail or a screw offer greater axial and rotational stability, the use of an intramedullary rod provides a sufficient stabilization for consistent healing outcomes without producing new defects in the bone tissue or damaging nearby soft tissue. Radiographic imaging is used to monitor the progression of callus formation, bony union, and subsequent remodeling of the bony callus. Bone healing outcomes are typically associated with the strength of the healed bone and measured with torsional testing. Still, understanding the early cellular and molecular events associated with fracture repair is critical in the study of bone tissue regeneration. The closed femoral fracture model in mice with intramedullary fixation serves as an attractive platform to study bone fracture healing and evaluate therapeutic strategies to accelerate healing.

The murine closed femoral fracture model is a powerful platform to study fracture healing and novel therapeutic strategies to accelerate bone regeneration. The goal of this surgical protocol is to generate unilateral closed femoral fractures in mice using an intramedullary steel rod to stabilize the femur.

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Conducting Maximal and Submaximal Endurance Exercise Testing to Measure Physiological and Biological Responses to Acute Exercise in Humans

Regular physical activity has a positive effect on human health, but the mechanisms controlling these effects remain unclear. The physiologic and biologic responses to acute exercise are predominantly influenced by the duration and intensity of the exercise regimen. As exercise is increasingly thought of as a therapeutic treatment and/or diagnostic tool, it is important that standardizable methodologies be utilized to understand the variability and to increase the reproducibility of exercise outputs and measurements of responses to such regimens. To that end, we describe two different cycling exercise regimens that yield different physiologic outputs. In a maximal exercise test, exercise intensity is continually increased with a greater workload resulting in an increasing cardiopulmonary and metabolic response (heart rate, stroke volume, ventilation, oxygen consumption and carbon dioxide production). In contrast, during endurance exercise tests, the demand is increased from that at rest, but is raised to a fixed submaximal exercise intensity resulting in a cardiopulmonary and metabolic response that typically plateaus. Along with the protocols, we provide suggestions on measuring physiologic outputs that include, but are not limited to, heart rate, slow and forced vital capacity, gas exchange metrics, and blood pressure to enable the comparison of exercise outputs between studies. Biospecimens can then be sampled to assess cellular, protein, and/or gene expression responses. Overall, this approach can be easily adapted into both short- and long-term effects of two distinct exercise regimens.

To assess the influence of exercise intensity on physiologic and biologic responses, two different exercise testing protocols were utilized. Methods outlining exercise testing on a cycle ergometer as an incremental maximal oxygen consumption test and endurance, steady state submaximal endurance test are described.

Regular physical activity has a positive effect on human health, but the mechanisms controlling these effects remain unclear. The physiologic and biologic ...

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Measurement of rodent skeletal muscle contractile function is a useful tool that can be used to track disease progression as well as efficacy of therapeutic intervention. We describe here the non-invasive, in vivo assessment of the dorsiflexor muscles that can be repeated over time in the same mouse.

Assessment of skeletal muscle contractile function is an important measurement for both clinical and research purposes. Numerous conditions can negatively ...

Implantation of an Isoproterenol Mini-Pump to Induce Heart Failure in Mice

Isoproterenol (ISO), is a non-selective beta-adrenergic agonist, that is used widely to induce cardiac injury in mice. While the acute model mimics stress-induced cardiomyopathy, the chronic model, administered through an osmotic pump, mimics advanced heart failure in humans. The purpose of the described protocol is to create the chronic ISO-induced heart failure model in mice using an implanted mini-pump. This protocol has been used to induce heart failure in 100+ strains of inbred mice. Techniques on surgical pump implantation are described in detail and may be relevant to anyone interested in creating a heart failure model in mice. In addition, the weekly cardiac remodeling changes based on echocardiographic parameters for each strain and expected time to model development are presented. In summary, the method is simple and reproducible. Continuous ISO administered via the implanted mini-pump over 3 to 4 weeks is sufficient to induce cardiac remodeling. Finally, the success for ISO model creation may be assessed in vivo by serial echocardiography demonstrating hypertrophy, ventricular dilation, and dysfunction.

The chronic administration of isoproterenol via an implanted osmotic pump has been used widely to mimic advanced heart failure in mice. Here, we describe detailed methods in surgical mini-pump implantation for the continuous isoproterenol administration over 3 weeks, as well as, echocardiographic assessment for the successful model creation.

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A Piglet Perinatal Asphyxia Model to Study Cardiac Injury and Hemodynamics after Cardiac Arrest, Resuscitation, and the Return of Spontaneous Circulation

Neonatal piglets have been extensively used as translational models for perinatal asphyxia. In 2007, we adapted a well-established piglet asphyxia model by introducing cardiac arrest. This enabled us to study the impact of severe asphyxia on key outcomes, including the time taken for the return of spontaneous circulation (ROSC), as well as the effect of chest compressions according to alternative protocols for cardiopulmonary resuscitation. Due to the anatomical and physiological similarities between piglets and human neonates, piglets serve as good models in studies of cardiopulmonary resuscitation and hemodynamic monitoring. In fact, this cardiac arrest model has provided evidence for guideline development through research on resuscitation protocols, pathophysiology, biomarkers, and novel methods for hemodynamic monitoring. Notably, the incidental finding that a substantial fraction of piglets have pulseless electrical activity (PEA) during cardiac arrest may increase the applicability of the model (i.e., it may be used to study pathophysiology extending beyond the perinatal period). However, the model generation is technically challenging and requires various skill sets, dedicated personnel, and a fine balance of the measures, including the surgical protocols and the use of sedatives/analgesics, to ensure a reasonable rate of survival. In this paper, the protocol is described in detail, as well as experiences with adaptations to the protocol over the years.

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Chronic Ovine Model of Right Ventricular Failure and Functional Tricuspid Regurgitation

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Right ventricular failure and functional tricuspid regurgitation are associated with left-sided heart disease and pulmonary hypertension, which contribute significantly to morbidity and mortality in patients. Establishing a chronic ovine model to study right ventricular failure and functional tricuspid regurgitation will help in understanding their mechanisms, progression, and possible treatments.

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Modified Heterotopic Abdominal Heart Transplantation and a Novel Aortic Regurgitation Model in Rats

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical technique. Modifying the transplantation procedure to strengthen the myocardial protection could prolong the ischemia time while preserving the donor&#39;s cardiac function. This technique&#39;s key points are as follows: transecting the donor&#39;s abdominal aorta before harvesting to unload the donor&#39;s heart; perfusing the donor&#39;s coronary arteries with a cold cardioplegic solution; and topical cooling of the donor&#39;s heart during the anastomosis procedure. Consequently, since this procedure prolongs the acceptable ischemia time, beginners can easily perform it and achieve a high success rate.
Moreover, a new aortic regurgitation (AR) model was established in this work using a technique different from the existing one,&#160;which is created by inserting a catheter from the right carotid artery and puncturing the native aortic valve under continuous echocardiographic guidance. A heterotopic abdominal heart transplantation was performed using the novel AR model. In the protocol, after the donor&#39;s heart is harvested, a stiff guidewire is inserted into the donor&#39;s brachiocephalic artery and advanced toward the aortic root. The aortic valve is punctured by pushing the guidewire further even after the resistance is felt, thus inducing AR. It is easier to damage the aortic valve using this method than with the procedure described in the conventional AR model. Additionally, this novel AR model does not contribute to the recipient&#39;s circulation; therefore, this method is expected to produce a more severe AR model than the conventional procedure.

This study demonstrates a reproducible heterotopic abdominal heart transplantation technique in rats that beginners can learn and perform. Additionally, a novel aortic regurgitation model in rats is generated by performing heterotopic abdominal heart transplantation and damaging the donor's aortic valve using a guidewire after harvesting.

Over the past 50 years, many researchers have reported heterotopic abdominal heart transplantation in mice and rats, with some variations in the surgical ...

In Vivo Heart Transplantation to Study Endothelial-to-Mesenchymal Transition

A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and precludes patients with congenital critical aortic stenosis and hypoplastic left heart syndrome (HLHS) from curative anatomical biventricular surgical repair. Surgical resection is currently the only available therapeutic option, but EFE often recurs, sometimes with an even more infiltrative growth pattern into the adjacent myocardium.
To better understand the underlying mechanisms of EFE and to explore therapeutic strategies, an animal model suitable for preclinical testing was developed. The animal model takes into consideration that EFE is a disease of the immature heart and is associated with flow disturbances, as supported by clinical observations. Thus, the heterotopic heart transplantation of neonatal rat donor hearts is the basis for this model.
A neonatal rat heart is transplanted into an adolescent rat&#39;s abdomen and connected to the recipient&#39;s infrarenal aorta and inferior vena cava. While perfusion of the coronary arteries preserves the viability of the donor heart, flow stagnation within the LV induces EFE growth in the very immature heart. The underlying mechanism of EFE formation is the transition of endocardial endothelial cells to mesenchymal cells (EndMT), which is a well-described mechanism of early embryonic development of the valves and septa but also the leading cause of fibrosis in heart failure. EFE formation can be macroscopically observed within days after transplantation. Transabdominal echocardiography is used to monitor the graft viability, contractility, and the patency of the anastomoses. Following euthanasia, the EFE tissue is harvested, and it shows the same histopathological characteristics as human EFE tissue from HLHS patients.
This in vivo model allows for studying the mechanisms of EFE development in the heart and testing treatment options to prevent this pathological tissue formation and provides the opportunity for a more generalized examination of EndMT-induced fibrosis.

Author Spotlight: A Neonatal Heterotopic Rat Heart Transplantation Model for the Study of Endothelial-to-Mesenchymal Transition  

This work presents an animal model of endothelial-to-mesenchymal transition-induced fibrosis, as seen in congenital cardiac defects such as critical aortic stenosis or hypoplastic left heart syndrome, which allows for detailed histological tissue evaluation, the identification of regulatory signaling pathways, and the testing of treatment options.

Endocardial fibroelastosis (EFE), defined by subendocardial tissue accumulation, has major impacts on the development of the left ventricle (LV) and ...