Special thanks to Chris Kassotis and Audrey Bailey for critical review of this manuscript and to Dr. Scott Korte, Joseph Beeman, Alison Curfman, Paul Kimball, Bridget Neibreggue, Jacob Redel, Amy Schroder, Maija Steinberg, and Stacey Winkeler for their assistance in optimization of this model in our laboratory. Funding was provided by the Clinical Biodetectives Training Grant (NIH T90) (KEP), University of Missouri Life Sciences Undergraduate Research Opportunities Program, MU Research Council, MU Research Board grants and NIH R21HD056441 (SCN).

1. Planning for live-animal surgery
<ol>
	<li>Ensure that appropriate approval has been received to work with laboratory animals.</li>
	<li>Order mice and allow at least one week of acclimation to their new environment.</li>
	<li>Female mice housed in the absence of exposure to male pheromones can stop cycling, a phenomenon referred to as the Whitten effect15,16. To keep mice cycling transfer urine-soaked male bedding to the female cage every five days. Alternatively, if open-top cages are used, place the female cage between two cages of males to keep the females cycling regularly.</li>
	<li>Ensure that mice are cycling by analyzing vaginal cytology daily for at least one week prior to surgery (Table 1)17.
	<ol>
		<li>Use a wax pencil to create eight partitions on a glass slide so that vaginal smears from multiple mice can be collected.</li>
		<li>Flush the vagina with 0.2-0.25 mL normal saline or distilled water using an eyedropper. Be sure to place the eyedropper just at the vaginal orifice, as cervical stimulation with the eyedropper could cause pseudopregnancy. Place the vaginal lavage on the glass slide for analysis of cell types. Slides can be read fresh (wet) or alternatively fixed by a number of methods and examined using a standard light microscope17.</li>
	</ol>
	</li>
	<li>Gather, clean and sterilize all necessary surgical equipment for a successful aseptic surgery (see Material Section)18.</li>
	<li>Prepare buprenorphine solution for analgesia in PBS using sterile technique to deliver 0.2 mg/kg final dose. The concentration of the buprenorphine solution should be 0.0333 mg/ml assuming that the average adult C57BL/6 mouse weighs approximately 0.025 kg and a subcutaneous injection volume of 0.15 ml per mouse. Buprenorphine can be prepared ahead of time and stored as aliquots. Note that buprenorphine is a Schedule III controlled substance requiring a DEA license and detailed inventory log.</li>
	<li>Prepare sterile PBS with penicillin (100 U/ml) and streptomycin (100 &mu;g/ml).</li>
	<li>Synchronize estrus cycles by transferring urine-soaked male bedding to the female cages 72 hours prior to induction15.</li>
</ol>
2. Prepare the surgical area for live animal surgery
<ol>
	<li>Prepare the surgical area as previously described18.</li>
	<li>Prepare the preparation area by setting out electric clippers, ophthalmic ointment, and surgical scrubs.</li>
	<li>Prepare the surgical area by placing a recirculating hot water heating pad on the surgical area to maintain body temperature throughout the surgery. Place a sterile waterproof pad over the recirculating hot water heating pad. Arrange surgical instruments, suture, sterile glass Petri dish, biopsy punch, sterile gauze, wound clips and wound clip applicator on the sterile surgical field.</li>
	<li>Prepare recovery area by placing recirculating hot water heating pads halfway under an empty cage to allow mice to move away from the heat if so desired.</li>
</ol>
3. Anesthetize and prepare the mouse for surgery
<ol>
	<li>Record the weight of the mouse and determine estrus stage by evaluating the vaginal cytology.</li>
	<li>For induction of anesthesia, place the mouse in an empty anesthesia chamber (empty cage with solid lid containing portal for isoflurane). Turn on isoflurane non-rebreathing anesthetic system and set the vaporizer to 4% isoflurane (with an oxygen flow rate of 0.5 to 1 L/min).</li>
	<li>When the mouse is under anesthesia switch the isoflurane flow to a cone (30-60 ml syringe sheath) and place mouse's nose and mouth in the cone on preparation table. Adequate anesthesia can be maintained with a lower concentration of isoflurane throughout the remainder of the surgery (~2.5-3.5% isoflurane). Adequate depth of anesthesia should be determined by a negative response to toe pinch stimulus.</li>
	<li>Apply ophthalmic ointment to avoid drying of the eyes during surgery.</li>
	<li>Using small electric clippers, shave the surgery site.</li>
	<li>Disinfect and prepare the surgery site with three alternating swipes of chlorhexidine scrub and 70% ethanol.</li>
	<li>Drape animal with a sterile field.</li>
</ol>
4. Uterine ligation
<ol>
	<li>Make a small (~1 cm) midline incision using either small scissors or a scalpel blade ending 0.5 - 1.0 cm rostral to the vaginal opening.</li>
	<li>Insert closed scissors into the opening such that the blades are between the body wall and abdominal wall. Gently blunt dissect the area around the incision by slowly opening and closing the scissors such that the abdominal wall is sufficiently detached from the skin. Remaining visible adhesions between the abdominal wall and skin around the incision site can be carefully snipped. Failure to adequately blunt dissect the incision site will make closing the abdominal wall more difficult.</li>
	<li>Using small forceps, gently locate the left uterine horn. The uterus is dorsal to the intestine, which is what you will see upon first entering the incision site. In some instances it is easiest to first locate the ovary and the associated ovarian fat pad. Gently pull up on the uterine horn and slide an open forceps underneath it to serve as a retractor. If desired, note the appearance of the ovaries and uterus at this time for additional information regarding the estrus stage at induction (Table 1).</li>
	<li>Gently slide two 6-8 cm pieces of 5-0 black braided silk suture (without a needle) underneath the stretched uterine horn.</li>
	<li>Securely ligate the horn at the utero-tubual junction (just caudal to the fallopian tube) and at the utero-cervical junction (just rostral to the cervix) using a square knot at each location. Leave the ends of the suture for the moment.</li>
	<li>Cut out the section of uterine horn between the two ligations and place the tissue in a sterile glass Petri dish containing ~100 &mu;L of PBS containing penicillin (100 U/ml) and streptomycin (100 &mu;g/ml). Cut the ends of the silk suture last. If suture comes loose or there is bleeding, find the stump and tie another knot.</li>
</ol>
5. Prepare endometriotic implants from excised uterus
<ol>
	<li>While the excised uterus is being manipulated, cover the abdomen with sterile gauze and maintain hydration with sterile PBS containing penicillin and streptomycin as needed.</li>
	<li>Strip the excised uterine horn of fat.</li>
	<li>If desired, weigh the excised uterine horn.</li>
	<li>Open the uterine horn by inserting one blade of small scissors (14 mm blade length) into the lumen and gently sliding the scissors down the uterine horn while holding the horn with forceps.</li>
	<li>In the glass Petri dish, use a 2 mm biopsy punch to cut out three equal sized implants.</li>
</ol>
6. Suturing endometriotic implants in peritoneal cavity
<ol>
	<li>Place sterile gauze immediately above the incision site and thoroughly wet with sterile PBS containing penicillin and streptomycin.</li>
	<li>With small, smooth forceps gently find the cecum and move rostrally along the small intestine. Pull out a small (4-5 cm) section of intestine that is at least two arteries away from the cecum and arrange it like a fan on the pre-wetted gauze so that the arterial cascade of the intestinal mesentery is clearly visible. Be sure to keep the bowel moist at all times with sterile saline. Note: do not use rat-toothed forceps while handling the bowel.</li>
	<li>Use 6-0 black ethilon suture with a P-1, 11 mm, 3/8 circle, reverse cutting needle to gently suture one implant to an artery approximately 0.5 cm from the bowel.</li>
	<li>Note: The intestinal mesentery is covered by a thin layer of peritoneum. Be careful to make a clean pass through this layer while suturing around the artery. Pull suture through slowly and carefully as not to tear the peritoneum or rupture the artery.</li>
	<li>Complete two knots of one throw each, being careful not to tighten the suture very hard, as doing so may result in loss of blood flow and subsequent necrosis of the intestine and death. Trim the suture within 2 mm of the implant. Wet bowel again to continue to maintain hydration before moving on to next implant.</li>
	<li>Moving in a rostral direction, pull out the next 3-4 cm of intestine and gently replace the section that already contains an implant. Skip one or two arteries from the previous implant site and suture the next implant. Repeat for the third implant.</li>
	<li>Replace all of the bowel in the abdominal cavity.</li>
</ol>
7. Sham surgeries
<ol>
	<li>Sham surgeries are performed using the same steps as the endometriosis surgeries except that no tissue is sutured to the intestinal mesentery.</li>
	<li>Excise the left uterine horn as in step 4.</li>
	<li>Endometriotic implants (step 5) are not prepared in the sham surgery. The excised uterine horn can be discarded or used for other purposes if desired.</li>
	<li>Sutures, but no tissues, are placed around three arteries in the arterial cascade of the intestinal mesentery as in step 6.</li>
</ol>
8. Closing the surgical wound
<ol>
	<li>Ensure that all of the organs are approximately back to their anatomical position.</li>
	<li>Use 5-0 coated vicryl suture in a non-interlocking continuous stitch to close the abdominal wall.</li>
	<li>Use 9 mm wound clips to close the skin.</li>
</ol>
9. Recover animal
<ol>
	<li>Administer 0.33 mg/ml buprenorphine at 0.15 ml/25 g mouse via subcutaneous injection for a dose of 0.2 mg/kg. Buprenorphine is administered post-operatively to prevent further cardiovascular/respiratory depression that can lengthen the recovery process.</li>
	<li>Gently dry the mouse with kimwipes or paper towels if it has gotten wet during the surgery.</li>
	<li>Place animal ventral side down in cage partially atop a recirculating heated water pad until the animal is recovered and has regained sternal recumbency (within five minutes as the inhalant anesthetic quickly wears off).</li>
</ol>
10. Post-operative care
<ol>
	<li>Mice should be observed every 15 minutes until they are able to maintain sternal recumbency, then hourly until they regain their normal behavior following surgery.</li>
	<li>Mice should appear normal within 24 hours of surgery. Mice should be monitored daily for seven to ten days for signs of recovery and good health.
	<ol>
		<li>Indications that an animal is in poor health, pain, or distress include decreased activity, self-mutilation, ungroomed appearance, or hunched posture.</li>
		<li>If an animal does not appear to be in good health within 24 hours of surgery, either administer buprenorphine (0.2 mg/kg) or euthanize the animal. If the animal does not improve within 8 hours of supplemental buprenorphine administration the animal should be euthanized as bowel necrosis is likely.</li>
	</ol>
	</li>
	<li>Remove wound clips 7-10 days post-induction.</li>
	<li>Continue to monitor estrus cyclicity by examination of vaginal cytology for the duration of the experiment. Synchronize estrus cycles 72 hours prior to collection by transferring urine-soaked male bedding to the female cages as described in step 1.3.</li>
</ol>
11. Necropsy and tissue excision
<ol>
	<li>The timing of necropsy is dependent on the particular research question and is further discussed in the representative results and discussion.</li>
	<li>Euthanize the mouse by carbon dioxide asphyxiation.</li>
	<li>Collect blood by cardiac puncture using a 23 gauge needle on a 1cc syringe (if desired).</li>
	<li>Collect a vaginal cytology smear as described above to determine estrus stage at the time of collection17.</li>
	<li>Cut remaining uterine horn at utero-tubal junction and at the cervix, remove fat, weigh, and process as desired (see 11.14 and 11.15).</li>
	<li>Locate the black sutures around the endometriotic lesions. Photograph intact endometriotic lesions if desired.</li>
	<li>Carefully dissect the adhesions surrounding the endometriotic lesions with a small scissors and forceps, being careful not to lance the lesions. Work quickly and carefully to prevent RNA degradation.</li>
	<li>Measure and record the length and width of the endometriotic lesions using calipers.</li>
	<li>Excise the endometriotic lesions and place on a paper towel moistened with PBS. Remove any non-endometriotic tissue from the lesions. A dissecting microscope or magnifying glass stand can be used to aid in the dissection.</li>
	<li>Weigh the three fluid filled endometriotic lesions prior to removing the suture.</li>
	<li>Gently remove the suture from the endometriotic lesions.</li>
	<li>For histology, formalin fix one fluid filled endometriotic lesion for two hours followed by three thirty minute PBS washes and final storage in 70% ethanol. Dehydrate and paraffin embed.</li>
	<li>Lance two of the endometriotic lesions. Weigh these again. Since cyclic hormonal changes can alter the amount of cyst fluid, this gives a measure of the tissue wet-weight in addition to the weight of the cyst plus the fluid as measured in 11.10.</li>
	<li>For RNA isolation and gene expression studies, immediately homogenize one of the lanced endometriotic lesions (or ~20 &mu;g uterine tissue) in lysis binding solution and store at -80&deg;C for future RNA isolation with the RNAqueous kit (Ambion) or other method as desired.</li>
	<li>For future isolation of RNA, DNA, or protein, immediately snap-freeze the second lanced endometriotic lesion (or ~20 &mu;g uterine tissue) in liquid nitrogen and store at -80&deg;C.</li>
</ol>
Representative Results
Endometriotic lesions in the mouse model of surgically induced endometriosis morphologically and histologically resemble those observed in women. Histological analysis of endometriosis in both women and the mouse model indicates that endometriotic lesions contain endometrial glands and stroma (Figure 2A). Endometriotic lesions in mice also contain hemosiderin-laden macrophages, which are a common hallmark of endometriosis in women (Figure 2B)19.
Endometriotic lesions removed from mice three days post-induction appear inflamed and hemorrhagic (Figure 3A). After two to four weeks of growth endometriotic lesions in the mouse model are cyst-like, fluid filled and surrounded by peritoneal adhesions (Figures 3B and 3C). Compared to lesion weight at induction, fluid filled lesions were 306% and 862% larger at one and two months post-induction and lanced lesions were 51% and 172% larger, respectively (Figures 4A and 4B). We have obtained consistent fluid filled and lanced endometriotic lesion weights at one-month post-induction over five different experiments (Figure 5). At one month post-induction fluid filled (7.44&plusmn;3.75 mg) and lanced (2.92&plusmn;1.23 mg) endometriotic lesion weight were significantly correlated (Pearson's correlation coefficient = 0.669, p &lt; 0.001).
Age of the mouse did not affect lesion size for mice between three and ten months of age. Neither the fluid filled or lanced endometriotic lesion weight at one month post-induction was significantly correlated with the age of the animal (r = -0.136, p = 0.380 and r = -0.063, p = 0.698, respectively).
The mouse uterus undergoes changes in size, fluid retention, cell proliferation and appearance due to the influence of steroid hormones during the estrus cycle. We compared the endometriotic lesion weight to the weight of the remaining intact uterine horn from animals in different estrus stages. We did not find a significant correlation between uterine weight and fluid filled or lanced endometriotic lesion weight at one-month post induction (r = -0.046, p = 0.765 and r = 0.232, p = 0.155, respectively).
The gene expression pattern observed in the endometriotic lesions of mice closely mirrors that reported in women with the disease5. By three days post-induction genes regulating extracellular matrix remodeling, cell adhesion, and angiogenesis are highly upregulated and many of these genes remain upregulated through one month of growth.
Figures and Tables
<img alt="Figure 1" src="/files/ftp_upload/3396/3396fig1.jpg" /> 
Figure 1. Surgical induction of endometriosis by autologus uterine tissue transfer in the mouse. The left uterine horn is ligated, excised, and opened longitudinally to expose the endometrium. Three 2 mm2 biopsies are prepared and each is sutured to an artery in the arterial cascade of the intestinal mesentery. By one month post-induction the endometriotic lesions are fluid filled and surrounded by adhesions.
<img alt="Figure 2" src="/files/ftp_upload/3396/3396fig2.jpg" /> 
Figure 2. Hematoxylin and eosin stained section of an endometrial lesion from the mouse model of endometriosis at one month post-induction demonstrating (A) the presence of endometrial glands and stroma; scale bar = 50 &mu;m and (B) hemosiderin-laden macrophages, some of which are indicated by arrows; scale bar = 20 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/3396/3396fig3.jpg" /> 
Figure 3. Endometriotic lesions in the mouse model following euthanasia, either three days post-induction (A) or one month post-induction (B and C).
<img alt="Figure 4" src="/files/ftp_upload/3396/3396fig4.jpg" /> 
Figure 4. Endometriotic lesions from mice surgically induced to have endometriosis were excised and weighed at one or two months post-induction. Data are average&plusmn;SEM. Data were log transformed and different letters indicate significance within each panel by one-way ANOVA followed by one-sided Fisher's Least Significant Difference Mulitple Comparisons. (A) Cyst like, fluid filled endometriotic lesions (N = 10, 7 or 5 for induction, one month, or two month post-induction, respectively). (B) Lanced endometriotic lesions (N = 10, 8 or 7 for induction, one month or two month post-induction respectively).
<img alt="Figure 5" src="/files/ftp_upload/3396/3396fig5.jpg" /> 
Figure 5. Endometriotic lesion wet weight with fluid and lanced at one month post-induction from five separate experiments. Data are average&plusmn;SEM. Mice N=10, 6, 8, 7 and 7 for fluid filled lesions and 0, 7, 10, 8, and 8 for lanced lesions in experiment 1, 2, 3, 4, and 5, respectively.
<img alt="Table 1" src="/files/ftp_upload/3396/3396table1.jpg" /> 
Table 1. Observation of Estrus Stage by Vaginal Cytology and Visual Appearance of Ovaries and Uterus and Induction. 
Appearance of ovary and uterus will be time dependent. The following are based on sacrifice around 8:00 am the morning of each cycle day. Further, the observations are subjective and comparing the ovary and uterine horns will be a better estimate than uterine horns only. These observations are meant to supplement the information obtained from daily vaginal cytology readings.
<img alt="Table 2" src="/files/ftp_upload/3396/3396table2.jpg" /> 
Table 2. Comparison of surgery in mouse and rat.

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+murine+endometriosis+induces+DNA+methylation+and+altered+gene+expression+in+eutopic+endometrium.">Experimental murine endometriosis induces DNA methylation and altered gene expression in eutopic endometrium.</a> Biol. Reprod. 80, 79-85 (2009).</li><li>Somigliana, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endometrial+ability+to+implant+in+ectopic+sites+can+be+prevented+by+interleukin-12+in+a+murine+model+of+endometriosis.">Endometrial ability to implant in ectopic sites can be prevented by interleukin-12 in a murine model of endometriosis.</a> Hum. Reprod. 14, 2944-2950 (1999).</li><li>Hirata, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+an+experimental+model+of+endometriosis+using+mice+that+ubiquitously+express+green+fluorescent+protein.">Development of an experimental model of endometriosis using mice that ubiquitously express green fluorescent protein.</a> Hum. Reprod. 20, 2092-2096 (2005).</li><li>Story, L., Kennedy, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+studies+in+endometriosis:+a+review.">Animal studies in endometriosis: a review.</a> Ilar. J. 45, 132-138 (2004).</li><li>Cummings, A. M., Hedge, J. M., Birnbaum, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+prenatal+exposure+to+TCDD+on+the+promotion+of+endometriotic+lesion+growth+by+TCDD+in+adult+female+rats+and+mice.">Effect of prenatal exposure to TCDD on the promotion of endometriotic lesion growth by TCDD in adult female rats and mice.</a> Toxicol. Sci. 52, 45-49 (1999).</li><li>Cummings, A. M., Metcalf, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+estrogen,+progesterone,+and+methoxychlor+on+surgically+induced+endometriosis+in+rats.">Effects of estrogen, progesterone, and methoxychlor on surgically induced endometriosis in rats.</a> Fundam. Appl. Toxicol. 27, 287-290 (1995).</li><li>Sharpe-Timms, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endometriotic+lesions+synthesize+and+secrete+a+haptoglobin-like+protein.">Endometriotic lesions synthesize and secrete a haptoglobin-like protein.</a> Biol. Reprod. 58, 988-994 (1998).</li><li>Yavuz, E., Oktem, M., Esinler, I., Toru, S. A., Zeyneloglu, H. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genistein+causes+regression+of+endometriotic+implants+in+the+rat+model.">Genistein causes regression of endometriotic implants in the rat model.</a> Fertil. Steril. 88, 1129-1134 (2007).</li><li>Dmitrieva, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endocannabinoid+involvement+in+endometriosis.">Endocannabinoid involvement in endometriosis.</a> Pain. 151, 703-710 (2010).</li><li>Efstathiou, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonsteroidal+antiinflammatory+drugs+differentially+suppress+endometriosis+in+a+murine+model.">Nonsteroidal antiinflammatory drugs differentially suppress endometriosis in a murine model.</a> Fertil. Steril. 83, 171-181 (2005).</li><li>Becker, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endostatin+inhibits+the+growth+of+endometriotic+lesions+but+does+not+affect+fertility.">Endostatin inhibits the growth of endometriotic lesions but does not affect fertility.</a> Fertil. Steril. 84, 1144-1155 (2005).</li><li>Becker, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Short+synthetic+endostatin+peptides+inhibit+endothelial+migration+in+vitro+and+endometriosis+in+a+mouse+model.">Short synthetic endostatin peptides inhibit endothelial migration in vitro and endometriosis in a mouse model.</a> Fertil. Steril. 85, 71-77 (2006).</li><li>Sharpe-Timms, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+rats+as+a+research+model+for+the+study+of+endometriosis.">Using rats as a research model for the study of endometriosis.</a> Ann. N.Y. Acad. Sci. 955, 318-327 (2002).</li><li>Stilley, J. A., Woods-Marshall, R., Sutovsky, M., Sutovsky, P., Sharpe-Timms, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reduced+Fecundity+in+Female+Rats+with+Surgically+Induced+Endometriosis+and+in+Their+Daughters:+A+Potential+Role+for+Tissue+Inhibitors+of+Metalloproteinase+1.">Reduced Fecundity in Female Rats with Surgically Induced Endometriosis and in Their Daughters: A Potential Role for Tissue Inhibitors of Metalloproteinase 1.</a> Biol. Reprod. 80, (2009).</li></ol>

No conflicts of interest declared.

There are several critical parameters that should be noted while performing the surgical induction of endometriosis in mice. First, endometriosis is an estrogen dependent disease and as such this surgery should be performed in intact animals or alternatively in ovariectomized animals supplemented with estrogens20. Second, suturing the endometrial biopsies to the arterial cascade must be performed with extreme care. We have found that using only two relatively loose knots with one throw each keeps the biopsy in...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent/equipment</td>
			<td>Company</td>
			<td>Catalogue number</td>
		</tr>
		<tr>
			<td>Wax pencil</td>
			<td>Fisher</td>
			<td>NC9954135</td>
		</tr>
		<tr>
			<td>Glass slide</td>
			<td>Fisher</td>
			<td>12-550-433</td>
		</tr>
		<tr>
			<td>Eyedropper</td>
			<td>Fisher</td>
			<td>S79383</td>
		</tr>
		<tr>
			<td>Standard light microscope for evaluating vaginal cytology smears</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Buprenorphine HCL c3 (CARJET) 10X1ml</td>
			<td>Butler Animal Health Supply</td>
			<td>022891</td>
		</tr>
		<tr>
			<td>Sterile phosphate buffered saline (PBS)</td>
			<td>Gibco</td>
			<td>14040-117</td>
		</tr>
		<tr>
			<td>10,000U/ml Penicillin, 10,000&mu;g/ml Streptomycin in 0.85% NaCl</td>
			<td>Hyclone</td>
			<td>SV30010</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Abbott Animal Health</td>
			<td>05260-05</td>
		</tr>
		<tr>
			<td>Isoflurane non-rebreathing anesthetic system</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Recirculating hot water heating pad</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>30 ml syringe sheath</td>
			<td>Fisher</td>
			<td>14-823-16G</td>
		</tr>
		<tr>
			<td>Powder free sterile gloves</td>
			<td>Fisherbrand</td>
			<td>19020558</td>
		</tr>
		<tr>
			<td>Ophthalmic ointment</td>
			<td>Major Pharmaceuticals</td>
			<td>10033691</td>
		</tr>
		<tr>
			<td>Small electrical clippers</td>
			<td>Wahl</td>
			<td>9861-600</td>
		</tr>
		<tr>
			<td>Chlorhexidine scrub</td>
			<td>Fisher</td>
			<td>NC9863042</td>
		</tr>
		<tr>
			<td>70% Ethanol</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Polylined sterile field</td>
			<td>Busse Hospital Disposables</td>
			<td>696</td>
		</tr>
		<tr>
			<td>Size 3 scalpel</td>
			<td>Fisher</td>
			<td>22-079-657</td>
		</tr>
		<tr>
			<td>Number 10 scalpel blades</td>
			<td>Fisher</td>
			<td>22-079-681</td>
		</tr>
		<tr>
			<td>Small surgical scissors</td>
			<td>Roboz</td>
			<td>RS-5850</td>
		</tr>
		<tr>
			<td>Small serrated semi-curved forceps</td>
			<td>Roboz</td>
			<td>RS-5135</td>
		</tr>
		<tr>
			<td>5-0 black braided silk suture</td>
			<td>Ethicon</td>
			<td>K870H</td>
		</tr>
		<tr>
			<td>Sterilized pyrex glass Petri dishes</td>
			<td>Corning</td>
			<td>70160-101</td>
		</tr>
		<tr>
			<td>2 mm biopsy punch</td>
			<td>Miltex</td>
			<td>33-31</td>
		</tr>
		<tr>
			<td>Sterile gauze</td>
			<td>Kendall</td>
			<td>1806</td>
		</tr>
		<tr>
			<td>6-0 black monofilament ethilon nylon suture</td>
			<td>Ethicon</td>
			<td>697G</td>
		</tr>
		<tr>
			<td>Needle drivers (optional)</td>
			<td>World Precision Instruments</td>
			<td>500023</td>
		</tr>
		<tr>
			<td>5-0 undyed braided coated vicryl suture</td>
			<td>Ethicon</td>
			<td>J490G</td>
		</tr>
		<tr>
			<td>9mm Autoclip wound clips</td>
			<td>Becton Dickinson</td>
			<td>427631</td>
		</tr>
		<tr>
			<td>Autoclip applier &amp; remover</td>
			<td>Becton Dickinson</td>
			<td>427630</td>
		</tr>
		<tr>
			<td>23G needle</td>
			<td>Becton Dickinson</td>
			<td>305193</td>
		</tr>
		<tr>
			<td>1cc syringe</td>
			<td>Becton Dickson</td>
			<td>301025</td>
		</tr>
		<tr>
			<td>5X magnifying glass stand (optional)</td>
			<td>Fisher</td>
			<td>14-648-23</td>
		</tr>
		<tr>
			<td>10% Buffered formalin</td>
			<td>Fisher</td>
			<td>SF100-4</td>
		</tr>
		<tr>
			<td>Calipers</td>
			<td>Roboz</td>
			<td>RS-6466</td>
		</tr>
		<tr>
			<td>Processing/embedding cassettes</td>
			<td>Fisher</td>
			<td>15-197-700A</td>
		</tr>
		<tr>
			<td>Biopsy foam pads</td>
			<td>Fisher</td>
			<td>22-038-222</td>
		</tr>
		<tr>
			<td>RNAqueous RNA isolation kit</td>
			<td>Ambion</td>
			<td>AM1912</td>
		</tr>
		<tr>
			<td>Liquid nitrogen</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Snap cap microcentrifuge flat top tube</td>
			<td>Fisher</td>
			<td>02-681-240</td>
		</tr>
		<tr>
			<td>Ketamine (optional)</td>
			<td>Simga</td>
			<td>K4138</td>
		</tr>
		<tr>
			<td>Domitor (medetomidine hydrochloride) (optional)</td>
			<td>Tocris</td>
			<td>2023</td>
		</tr>
		<tr>
			<td>Antisedan (atipamezole) (optional)</td>
			<td>Sigma</td>
			<td>A9611</td>
		</tr>
	</tbody>
</table>

mouse model of surgically-induced endometriosis by auto-transplantation of uterine tissue

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of lesions, and/or chronic pharmaceutical management of pain and infertility symptoms. The cost associated with endometriosis has been estimated at 22 billion dollars per year in the United States1. To further our understanding of mechanisms underlying this enigmatic disease, animal models have been employed. Primates spontaneously develop endometriosis and therefore primate models most closely resemble the disease in women. Rodent models, however, are more cost effective and readily available2. The model that we describe here involves an autologous transfer of uterine tissue to the intestinal mesentery (Figure 1) and was first developed in the rat3 and later transferred to the mouse4. The goal of the autologous rodent model of surgically-induced endometriosis is to mimic the disease in women. We and others have previously shown that the altered gene expression pattern observed in endometriotic lesions from mice or rats mirrors that observed in women with the disease5,6. One advantage of performing the surgery in the mouse is that the abundance of transgenic mouse strains available can aid researchers in determining the role of specific components important in the establishment and growth of endometriosis. An alternative model in which excised human endometrial fragments are introduced to the peritoneum of immunocompromised mice is also widely used but is limited by the lack of a normal immune system which is thought to be important in endometriosis2,7. Importantly, the mouse model of surgically induced endometriosis is a versatile model that has been used to study how the immune system8, hormones9,10 and environmental factors11,12 affect endometriosis as well as the effects of endometriosis on fertility13 and pain14.

Watch this Scientific Journal Video about Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue at JoVE.com

Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue

A description of the surgical induction of endometriosis in mice and rats by auto-transplantation of uterine tissue to the arterial cascade of the intestinal mesentery.

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of ...

mouse-model-surgically-induced-endometriosis-auto-transplantation

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28.4K Views.  University of Missouri. A description of the surgical induction of endometriosis in mice and rats by auto-transplantation of uterine tissue to the arterial cascade of the intestinal mesentery.

Video: Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue

Mouse Model of Intraluminal MCAO: Cerebral Infarct Evaluation by Cresyl Violet Staining

Stroke is the third cause of mortality and the leading cause of disability in the World. Ischemic stroke accounts for approximately 80% of all strokes. However, the thrombolytic tissue plasminogen activator (tPA) is the only treatment of acute ischemic stroke that exists. This led researchers to develop several ischemic stroke models in a variety of species. Two major types of rodent models have been developed: models of global cerebral ischemia or focal cerebral ischemia. To mimic ischemic stroke in patients, in whom approximately 80% thrombotic or embolic strokes occur in the territory of the middle cerebral artery (MCA), the intraluminal middle cerebral artery occlusion (MCAO) model is quite relevant for stroke studies. This model was first developed in rats by Koizumi et al. in 1986 1. Because of the ease of genetic manipulation in mice, these models have also been developed in this species 2-3.
	Herein, we present the transient MCA occlusion procedure in C57/Bl6 mice. Previous studies have reported that physical properties of the occluder such as tip diameter, length, shape, and flexibility are critical for the reproducibility of the infarct volume 4. Herein, a commercial silicon coated monofilaments (Doccol Corporation) have been used. Another great advantage is that this monofilament reduces the risk to induce subarachnoid hemorrhages. Using the Zeiss stereo-microscope Stemi 2000, the silicon coated monofilament was introduced into the internal carotid artery (ICA) via a cut in the external carotid artery (ECA) until the monofilament occludes the base of the MCA. Blood flow was restored 1 hour later by removal of the monofilament to mimic the restoration of blood flow after lysis of a thromboembolic clot in humans. The extent of cerebral infarct may be evaluated first by a neurologic score and by the measurement of the infarct volume. Ischemic mice were thus analyzed for their neurologic score at different post-reperfusion times. To evaluate the infarct volume, staining with 2,3,5-triphenyltetrazolium chloride (TTC) was usually performed. Herein, we used cresyl violet staining since it offers the opportunity to test many critical markers by immunohistochemistry. In this video, we report the MCAO procedure; neurological scores and the evaluation of the infarct volume by cresyl violet staining.

The intraluminal middle cerebral occlusion model in mice is herein presented. The extent of cerebral infarct is evaluated by a neurologic score and cresyl violet staining, an alternative staining to TTC, offering the great advantage to test in parallel many interest markers.

Stroke is the third  cause of mortality and the leading cause of disability in the World. Ischemic  stroke accounts for approximately 80% of all strokes. ...

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

Implantation of Inferior Vena Cava Interposition Graft in Mouse Model

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. The long-term clinical results showed excellent patency rates, however, with significant incidence of stenosis. To investigate the cellular and molecular mechanisms of vascular neotissue formation and prevent stenosis development in tissue engineered vascular grafts (TEVGs), we developed a mouse model of the graft with approximately 1 mm internal diameter. First, the TEVGs were assembled from biodegradable tubular scaffolds fabricated from a polyglycolic acid nonwoven felt mesh coated with &#949;-caprolactone and L-lactide copolymer. The scaffolds were then placed in a lyophilizer, vacuumed for 24 hr, and stored in a desiccator until cell seeding. Second, bone marrow was collected from donor mice and mononuclear cells were isolated by density gradient centrifugation. Third, approximately one million cells were seeded on a scaffold and incubated O/N. Finally, the seeded scaffolds were then implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. The implanted grafts demonstrated excellent patency (&#62;90%) without evidence of thromboembolic complications or aneurysmal formation. This murine model will aid us in understanding and quantifying the cellular and molecular mechanisms of neotissue formation in the TEVG.

To improve our knowledge of cellular and molecular neotissue formation, a murine model of the TEVG was recently developed. The grafts were implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. This model achieves similar results to those achieved in our clinical investigation, but over a far shortened time-course.

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. ...

Transplantation of Pulmonary Valve Using a Mouse Model of Heterotopic Heart Transplantation

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed results. However, the cellular and molecular mechanisms of the neotissue development, valve thickening, and stenosis development are not researched extensively. To answer the above questions, we developed a murine heterotopic heart valve transplantation model. A heart valve was harvested from a valve donor mouse and transplanted to a heart donor mouse. The heart with a new valve was transplanted heterotopically to a recipient mouse. The transplanted heart showed its own heartbeat, independent of the recipient&#8217;s heartbeat. The blood flow was quantified using a high frequency ultrasound system with a pulsed wave Doppler. The flow through the implanted pulmonary valve showed forward flow with minimal regurgitation and the peak flow was close to 100 mm/sec. This murine model of heart valve transplantation is highly versatile, so it can be modified and adapted to provide different hemodynamic environments and/or can be used with various transgenic mice to study neotissue development in a tissue engineered heart valve.

In order to understand the cellular and molecular mechanisms underlying neotissue formation and stenosis development in tissue engineered heart valves, a murine model of heterotopic heart valve transplantation was developed. A pulmonary heart valve was transplanted to recipient using the heterotopic heart transplantation technique.

Tissue engineered heart valves, especially decellularized valves, are starting to gain momentum in clinical use of reconstructive surgery with mixed ...

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo: A Model of Single Vessel Stroke

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to image cellular events in an intact animal. Additionally, the ability to induce physiological disease states such as stroke in vivo increases its utility. The technique described herein allows for physiological assessment of cellular responses within the CNS following a stroke and can be adapted for other pathological conditions being studied. The technique presented uses laser excitation of the photosensitive dye Rose Bengal in vivo to induce a focal ischemic event in a single blood vessel. 
The video protocol demonstrates the preparation of a thin-skulled cranial window over the somatosensory cortex in a mouse for the induction of a Rose Bengal photothrombotic event keeping injury to the underlying dura matter and brain at a minimum. Surgical preparation is initially performed under a dissecting microscope with a custom-made surgical/imaging platform, which is then transferred to a confocal microscope equipped with an inverted objective adaptor. Representative images acquired utilizing this protocol are presented as well as time-lapse sequences of stroke induction. This technique is powerful in that the same area can be imaged repeatedly on subsequent days facilitating longitudinal in vivo studies of pathological processes following stroke.

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo:  A Model of Single Vessel Stroke

Here, we describe a semi-invasive optical microscopy approach for the induction of a Rose Bengal photothrombotic clot in the somatosensory cortex of a mouse in vivo. The technical aspects of the imaging procedure are described from induction of a photothrombotic event to application and data collection.

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to ...

Creation of a Rodent Model of Abdominal Aortic Aneurysm by Blocking Adventitial Vasa Vasorum Perfusion

The adventitial vasa vasorum (VV) provides oxygen and nourishment to the aortic wall. Hypoxia in the aortic wall can cause enlarged abdominal aortic aneurysms (AAAs). This article introduces and describes a standard protocol used to induce AAAs through adventitial VV hypoperfusion created with a combination of polyurethane catheter insertion into the aortic lumen and suture ligation of the infrarenal abdominal aorta.
The protocol involves the use of male rats weighing 300-400 g, which are provided food and water ad libitum. After laparotomy with a ventral midline abdominal incision, exfoliation of the aorta is performed, which blocks blood flow from the perivascular tissue. Aortotomy involving a small incision adjacent to the renal artery branches is performed, and a polyurethane catheter is inserted using an 18-gauge indwelling needle. After repairing the incision, tight ligation of the aorta over the catheter blocks VV blood flow from the proximal direction through the aortic wall without disturbing the aortic blood flow. This technique can induce an AAA with progressive aortic dilatation.
The greatest benefit of this model is that VV hypoperfusion causes tissue hypoxia and the development of an infrarenal AAA, which has morphological and pathological characteristics similar to those of a human AAA.

Polyurethane catheter insertion into the aortic lumen and suture ligation of the aorta induce chronic hypoxia due to hypoperfusion of the adventitial vasa vasorum. This article describes a novel animal model of abdominal aortic aneurysm (AAA) with characteristics similar to those of AAA in humans.

The adventitial vasa vasorum (VV) provides oxygen and nourishment to the aortic wall. Hypoxia in the aortic wall can cause enlarged abdominal aortic ...

A Mouse Model of Orthopedic Surgery to Study Postoperative Cognitive Dysfunction and Tissue Regeneration

Surgery is commonly used to improve and maintain quality of life. Unfortunately, in vulnerable patients such as the elderly, complications may occur and significantly diminish the outcome. Indeed, after routine orthopedic surgery to repair a fracture, as many as 50% of elderly patients suffer from neurologic complications like delirium. Also, the capacity to heal and regenerate tissue after surgery decreases with age, and can impact the quality of fracture repair and even osseous integration of implants. Thus, a better understanding of mechanisms that drive these age-dependent changes could provide strategic targets to minimize risk for such complications and optimize outcomes. Here, we introduce a clinically relevant mouse model of tibial fracture. The postoperative changes in these mice mimic some of the cognitive impairments commonly observed after routine orthopedic surgery in humans. Briefly, an incision is performed in the right hind limb under strictly aseptic conditions. Muscles are disassociated, and a 0.38-mm stainless steel pin is inserted into the upper crest of the tibia, inside the intramedullary canal. Osteotomy is then performed, and the wound is stapled. We have used this model to investigate the effects of surgical trauma on postoperative neuroinflammation and behavioral changes. By applying this fracture model in combination with parabiosis, a surgical model in which 2 mice are anastomosed, we have studied cells and secreted factors that systemically rejuvenate organ function and tissue regeneration after injury. By following our step-by-step protocol, these models can be reproduced with high fidelity, and can be adapted to interrogate many biologic pathways that are altered by surgical trauma.

This protocol describes a mouse model of orthopedic surgery that has been used to study mechanisms of postoperative neuroinflammation and behavioral changes, and when combined with parabiosis, to study tissue regeneration during aging.

Surgery is commonly used to improve and maintain quality of life. Unfortunately, in vulnerable patients such as the elderly, complications may occur and ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing (Neo)adjuvant Therapies

Surgery is often the first treatment for many solid tumors. However, local relapses frequently occur following primary tumor resection, despite adjuvant or neo-adjuvant therapies. This occurs when surgical margins are insufficiently tumor-free, resulting in residual cancer cells. From a biological and immunological perspective, surgery is not a null event; the wound healing environment is known to induce both pro- and anti-tumorigenic pathways. As a consequence, preclinical models for drug development aimed at preventing local relapse should incorporate surgical resection when testing new (neo)adjuvant therapies, to model the clinical settings in patients treated with surgery.
Here, we describe a mouse model of incomplete surgical resection of WEHI 164 soft tissue sarcoma that allows testing of (neo)adjuvant therapies in the setting of a wound healing response. In this model, 50% or 75% of the tumor is removed, leaving behind some cancer tissue in situ to model gross residual disease after surgery in the clinical setting. This model allows testing therapies in the context of surgery while also considering the wound healing response, which may affect the efficacy of (neo)adjuvant treatments. The incomplete surgical resection results in reproducible regrowth of the tumor in all mice in the absence of adjuvant therapy. Adjuvant treatment with checkpoint blockade results in reduced tumor regrowth. This model is thus appropriate for testing therapies in the context of debulking surgery and its associated wound healing response and can be extended to other types of solid cancer.

A Mouse Model of Incompletely Resected Soft Tissue Sarcoma for Testing Neoadjuvant Therapies

In this protocol, we describe a mouse model of incomplete surgical resection of soft tissue sarcoma for testing (neo)adjuvant therapies.

Surgery is often the first treatment for many solid tumors. However, local relapses frequently occur following primary tumor resection, despite adjuvant ...