We thank members of the Mariani lab for critical reading of the manuscript. Our funding sources were: the Baxter Medical Scholar Research Fellowship (to M.K.S.), USC undergraduate fellowships and the Provost, Dean Joan M. Schaeffer, and Rose Hills Fellowships, (to M.K.S.). We also acknowledge a CIRM BRIDGES fellowship through Pasadena City College (to T.T.T). and the James H. Zumberge Research and Innovation Fund, the USC Regenerative Medicine Initiative, and the NIAMS NIH under Award Number R21AR064462 (to F.V.M).

NOTE: All procedures are in accordance with an animal protocol approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Southern California.
1. Preparation for Surgery
<ol>
	<li>Lay out the necessary surgical tools and supplies (sterilized, items 1 - 18 in Materials list) and arrange a dissecting microscope and a fiber optics trans-illuminator in a hood or on a downdraft table.</li>
	<li>Place a sexually mature mouse (20 - 30 g, 1 - 3 months old) in an induction chamber with 4% isoflurane to induce anesthesia.</li>
	<li>Confirm anesthesia with a paw and/or tail pinch test.</li>
	<li>Apply ointment to the mouse&#8217;s eyes to avoid eye dryness during the surgery.</li>
	<li>Move the mouse to the microscope stage. Place the mouse into the nose cone of the maintenance hose and position laterally. Provide a warming pouch adjacent to the animal to help maintain body temperature. 
	NOTE: Position of the mouse is dependent on personal preference and/or handedness of the surgeon., either left or right side is fine.</li>
	<li>Adjust isoflurane to 2 - 3% for maintenance, and watch breathing rate.</li>
	<li>Inject buprenorphine (0.05 mg/kg) subcutaneously (lateral flank of the leg) for post-surgery pain with a 25 G needle. The leg may twitch which is a normal response.</li>
</ol>
2. Opening Incision
<ol>
	<li>Detect the desired rib by palpation, and clear the area of hair with a shaver. 
	NOTE: Ribs 8-10 are recommended since ribs 1-7 are closer to the lungs and 11-13 are considerably shorter. Counting the ribs should begin with the most rostral rib as number 1.</li>
	<li>Prepare the area with alternating povidone-iodine and isopropanol (70%) swabbing (3x is often recommended by animal use protocols).</li>
	<li>Cut a 2 cm incision through the skin, directly above and parallel to the desired rib with medium sized microsurgery scissors. Incise through the underlying muscle and fat layers.</li>
	<li>Place all 3 layers (skin, muscle, fat) into a retractor to expose the surgical area, while minimizing the size of the incision.</li>
</ol>
3. Excising the Rib
<ol>
	<li>Cut through the intercostal muscles overlying the desired section of rib with a 5.0 mm scalpel. Position this incision around 5 mm proximal to the chondrocostal joint as this is where the rib is not as highly curved. Carefully separate the muscle from the bone with fine tip forceps.</li>
	<li>To create a resection while retaining the periosteum in the animal, cut through the periosteum, along the length of rib with a 5.0 mm scalpel. Carefully separate the periosteum from the underlying bone laterally with fine tip forceps. Proceed with caution, as the periosteum is very delicate and has a gelatinous consistency.</li>
	<li>Next make a cross-section through the bone at one end with fine micro-scissors. If needed, measure the resection with a reticle in the microscope or a ruler type gauge. Then carefully lift the bone out of the periosteum and cut the other end. 
	NOTE: Proceed with caution. This is the most delicate step, as pulling up without excising the rib below will tear the pleural membrane and result in a pneumothorax. If the pleural membrane is torn this will be readily evident as the interior of the thoracic cavity will become visible.</li>
	<li>If bleeding occurs as a result of cutting the bone, apply pressure at the cut end with a cotton-tipped swab for 4 - 5 sec to stop the bleeding. 
	NOTE: On the rare occasion that bleeding continues, it may be necessary to stop and euthanize the animal as significant blood loss will compromise recovery (without fluid replacement, the maximum blood volume which can be safely be lost is 10% of the total blood volume or 7.7 - 8 &#181;l/g. For a 25 g mouse, this is equivalent to around 180 - 200 &#181;l9).</li>
	<li>Immediately place the removed rib into 4% PFA for future analysis. 
	NOTE: If the resection of both the bone and the periosteum is to be carried out, omit the periosteum incision and separation (step 3.2). Since the periosteum is very tightly attached to the pleural membrane proceed cautiously, teasing the rib away from the pleural membrane with forceps to avoid tearing.</li>
</ol>
4. Closing the Incision
<ol>
	<li>Suture the intercostal muscles over the top of the remaining periosteal sleeve with 9-0 sutures (2 sutures are usually sufficient). Place the sutures directly above the cut ends of the rib, to act as indicators for surgery location.</li>
	<li>Remove the retractor. Suture the overlying muscle and fat with 9-0 sutures (3 - 4 sutures are usually sufficient). In mice with particularly thick layers of muscle and/or fat, suture each layer separately (e.g., 1 layer of sutures for the muscular layer and 1 layer of sutures for the fatty layer).</li>
	<li>Close the skin with 7-0 sutures (4 - 5 sutures are usually sufficient).</li>
	<li>Secure the incision with suture glue, pinching the edges together with large forceps.</li>
	<li>Slowly wean the mouse off isoflurane by first adjusting to 1% for few minutes and then turn off.</li>
	<li>Place the mouse under a heat lamp and leave until consciousness is regained, 5 - 10 min. After consciousness is regained and throughout the healing period, the mouse should move and ambulate normally and show no signs of distress.</li>
</ol>
5. Recovery and Analysis
<ol>
	<li>Provide post-operative pain management including the administration of buprenorphine in an oral gelatin form at 0.5 mg/kg every 12 hr for 48 hr. Oral administration does not require restraining the animal which might cause pain to the thoracic region.</li>
	<li>Maintain the mouse in a cage with free access to food and water during the healing period. After the incision is dry, female animals can be co-housed while males must remain isolated to prevent fighting.</li>
	<li>After the healing period, follow the approved procedure for euthanasia. Remove the rib cage for fixation and analysis.</li>
</ol>

<ol>
	<li>Philip, S. J., Kumar, R. J., Menon, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+study+of+rib+regeneration+following+costectomy+in+adolescent+idiopathic+scoliosis.">Morphological study of rib regeneration following costectomy in adolescent idiopathic scoliosis.</a> Eur Spine J. 14 (8), 772-776 (2005).</li><li>Munro, I. R., Guyuron, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Split-rib+cranioplasty.">Split-rib cranioplasty.</a> Ann Plast Surg. 7 (5), 341-346 (1981).</li><li>Taggard, D. A., Menezes, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+use+of+rib+grafts+for+cranioplasty+in+children.+Pediatric+neurosurgery.">Successful use of rib grafts for cranioplasty in children. Pediatric neurosurgery.</a> 34 (3), 149-155 (2001).</li><li>Head, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prevention+of+Regeneration+fo+the+Ribs:+A+problem+in+thoracic+surgery.">Prevention of Regeneration fo the Ribs: A problem in thoracic surgery.</a> Archives of Surgery. 14 (6), 1215-1221 (1927).</li><li>Kawanabe, Y., Nagata, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+of+costal+cartilage+harvest+for+total+auricular+reconstruction:+part+I.+Avoidance+and+prevention+of+intraoperative+and+postoperative+complications+and+problems.">A new method of costal cartilage harvest for total auricular reconstruction: part I. Avoidance and prevention of intraoperative and postoperative complications and problems.</a> Plastic and reconstructive surgery. 117 (6), 2011-2018 (2006).</li><li>Cheung, K. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+externally+fixed+femoral+fracture+model+for+mice.">An externally fixed femoral fracture model for mice.</a> Journal of orthopaedic research : official publication of the Orthopaedic Research Society. 21 (4), 685-690 (2003).</li><li>Matthys, R., Perren, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Internal+fixator+for+use+in+the+mouse.">Internal fixator for use in the mouse.</a> Injury. 40, S103-S109 (2009).</li><li>Cooper, G. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Testing+the+critical+size+in+calvarial+bone+defects:+revisiting+the+concept+of+a+critical-size+defect.">Testing the critical size in calvarial bone defects: revisiting the concept of a critical-size defect.</a> Plastic and reconstructive surgery. 125 (6), 1685-1692 (2010).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ask+the+Vet.">Ask the Vet.</a> JAX NOTES. 499, (2005).</li><li>Flecknell, P. A., Roughan, J. V., Stewart, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+oral+buprenorphine+('buprenorphine+jello')+for+postoperative+analgesia+in+rats--a+clinical+trial.">Use of oral buprenorphine ('buprenorphine jello') for postoperative analgesia in rats--a clinical trial.</a> Laboratory animals. 33 (2), 169-174 (1999).</li><li>Rigueur, D., Lyons, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-mount+skeletal+staining.">Whole-mount skeletal staining.</a> Methods in molecular biology. 1130, 113-121 (2014).</li><li>Evans, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contribution+of+somitic+cells+to+the+avian+ribs.">Contribution of somitic cells to the avian ribs.</a> Developmental biology. 256 (1), 114-126 (2003).</li><li>Colnot, C., Thompson, Z., Miclau, T., Werb, Z., Helms, 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=Altered+fracture+repair+in+the+absence+of+MMP9.">Altered fracture repair in the absence of MMP9.</a> Development. 130 (17), 4123-4133 (2003).</li><li>Lu, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+basis+for+age-related+changes+in+fracture+repair.">Cellular basis for age-related changes in fracture repair.</a> Journal of orthopaedic research : official publication of the Orthopaedic Research Society. 23 (6), 1300-1307 (2005).</li><li>Zilberman, Y., Gafni, Y., Pelled, G., Gazit, Z., Gazit, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioluminescent+imaging+in+bone.">Bioluminescent imaging in bone.</a> Methods in molecular biology. 455, 261-272 (2008).</li><li>Pelled, G., Gazit, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+using+osteocalcin-luciferase.">Imaging using osteocalcin-luciferase.</a> Journal of musculoskeletal. 4 (4), 362-363 (2004).</li><li>Elefteriou, F., Yang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+mouse+models+for+bone+studies--strengths+and+limitations.">Genetic mouse models for bone studies--strengths and limitations.</a> Bone. 49 (6), 1242-1254 (2011).</li><li>Srour, M. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Natural+large-scale+regeneration+of+rib+cartilage+in+a+mouse.">Natural large-scale regeneration of rib cartilage in a mouse.</a> J. Bone Miner. , (2014).</li></ol>

The authors have nothing to disclose

When first learning this protocol, determining where to locate the initial incision can be challenging. However, practice on euthanized mice helps the surgeon learn where to place the initial incision and expose the desired rib to be resected. Working on cadavers also improves the fine-motor skills required to remove the rib portion with or without the periosteum. In addition, someone new to this procedure might find manipulating the fine tools and thin sutures to be difficult. While tying off, excess pulling on the sutu...

Debilitating skeletal injury, chronic osteoarthritis, and the severe problems associated with reconstructive surgery impact economic productivity, family well-being, and quality of life. While small breaks and lesions can heal fairly well, humans are not capable of repairing large defects and therefore must rely on reconstructive procedures&#160;to restore structure and function. Reconstruction may involve allogeneic or heterogeneic grafts, morcellized bone, implanted scaffolds, or distraction osteogenesis. Unfortunately, not only are there persistant morbidity factors associated with these treatments but the original strength of the repaired bone is rarely attained. Thus, new clinical approaches are needed.
One-way to develop innovative methods to treat segmental defects is to study situations in which large-scale repair occurs naturally. Amphibians famously can regenerate skeletal elements, while mammals are considered limited in this ability. However, since the early part of the 20th century, a few reports of regeneration in the human rib have been published suggesting that humans may not be so limited1-4 . Currently this phenomenon is best known by plastic surgeons who use rib material for jaw, face and ear reconstruction, but it is not more broadly appreciated5. In order to study this repair in more detail, we have developed a surgical model using the mouse. Using this protocol, researchers can identify the innate factors involved and use this information to facilitate skeletal healing in other locations.
There are many advantages to using the ribs as a model for studying skeletal repair. First, the surrounding ribs provide a natural fixator (as compared to resection of the femur6,7). This decreases the morbidity risk of internal and external fixators and simplifies the surgical procedure. Secondly, the thin muscular layers of the chest wall provide for easy access and excellent visibility which make the assay comparable to the convenience of calvarial resections8. Thirdly, in contrast to the calvariae which form by intramembranous ossification, the ribs form by endochondral ossification and grow in length via extension at growth plates located at either end of a central diaphysis. Therefore, repair of the ribs may be more comparable to repair of the long bones of the appendicular skeleton. Furthermore, we have found that compared to the femur, the periosteum of the rib is thicker and can be more easily manipulated. Thus, investigators who wish to assay bone repair for the purpose of studying the periosteum or testing cell therapies, pharmacological agents, and/or tissue scaffolds may find this surgical model useful. In summary, this rib resection model provides a context within which to study natural large-scale bone repair in mammals as no such model in general use currently exists.

<table border="0" cellpadding="0" cellspacing="0" width="896">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Name of Reagent/ Equipment</td>
			<td style="width:160px;">Company</td>
			<td style="width:157px;">Catalog Number</td>
			<td style="width:304px;">Comments/Description</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:275px;">Medium sized micro-dissection scissors (Vannas-T&#252;bingen Spring Scissors 5 mm)</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td>15003-08</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">Fine micro-dissection scissors (Vannas Spring Scissors - 2mm Cutting Edge)</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td style="width:157px;">15000-04</td>
			<td>curved tip is beneficial</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Micro-scalpel 5.0 mm</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td>10315-12</td>
			<td>other fine scalpels can be substituted</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Dumont 55 forceps</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td style="width:157px;">11295-51</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Retractor&#160;</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td>17004-05</td>
			<td>adjustability is convenient</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Micro-needle holders</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td style="width:157px;">12060-01</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">9.0 nylon sutures (Ethilon), taper point best</td>
			<td style="width:160px;">Ethicon</td>
			<td>2819G or similar</td>
			<td>taper point best but reverse cutting is also good</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">7.0 prolene sutures (Prolene)</td>
			<td style="width:160px;">Ethicon</td>
			<td>8700H or similar</td>
			<td>6-0 can be used too, needle point can vary</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Large forceps (Adson Forceps)</td>
			<td style="width:160px;">Fine Science Tools</td>
			<td style="width:157px;">11006-12</td>
			<td>other brands are fine</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Lubricant Eye Ointment (Akwa Tears)</td>
			<td style="width:160px;">Akorn</td>
			<td style="width:157px;">17478-062-35</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:275px;">Suture glue (GLUture Topical Tissue Adhesive)</td>
			<td style="width:160px;">Abbot</td>
			<td style="width:157px;">32046-01</td>
			<td>has excellent working time</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Shaver</td>
			<td style="width:160px;">Wahl</td>
			<td>9918-6171 or similar</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Clamp lamp</td>
			<td>Zoo Med</td>
			<td style="width:157px;">LF-5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Infrared Bulb, 75W</td>
			<td style="width:160px;">Zoo Med</td>
			<td>RS-75</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:275px;">RC2 Rodent Anesthesia System</td>
			<td style="width:160px;">VetEquip&#160;</td>
			<td style="width:157px;">922100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">IsoFlo (Isoflurane)</td>
			<td style="width:160px;">Abbot</td>
			<td style="width:157px;">05260-05</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Buprenorphine (Buprenex)</td>
			<td style="width:160px;">Reckitt Benckiser</td>
			<td style="width:157px;">12496-0757-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Betadine</td>
			<td style="width:160px;">Purdue Frederick</td>
			<td>67618015017</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:275px;">Flavored Gelatin, raspberry</td>
			<td style="width:160px;">Jell-O</td>
			<td>B000E1FYL0</td>
			<td>made up firm, to the consistency of &#39;jigglers&#39;</td>
		</tr>
	</tbody>
</table>

a surgical procedure for resecting the mouse rib: a model for large-scale long bone repair

This protocol introduces researchers to a new model for large-scale bone repair utilizing the mouse rib. The procedure details the following: preparation of the animal for surgery, opening the thoracic body wall, exposing the desired rib from the surrounding intercostal muscles, excising the desired section of rib without inducing a pneumothorax, and closing the incisions. Compared to the bones of the appendicular skeleton, the ribs are highly accessible. In addition, no internal or external fixator is necessary since the adjacent ribs provide a natural fixation. The surgery uses commercially available supplies, is straightforward to learn, and well-tolerated by the animal. The procedure can be carried out with or without removing the surrounding periosteum, and therefore the contribution of the periosteum to repair can be assessed. Results indicate that if the periosteum is retained, robust repair occurs in 1 - 2 months. We expect that use of this protocol will stimulate research into rib repair and that the findings will facilitate the development of new ways to stimulate bone repair in other locations around the body.

Animals typically recover quickly from this procedure, the incision heals well, and the animals exhibit normal behavior. The procedure uses commercially available supplies and after practice, can be completed in under 1 hr. Before using this procedure to generate experimental data, it is important to analyze the resection at the day 0 time point to determine if any large bone fragments might remain within the resection zone. One way to assess proper procedure is to assess the animals by X-ray or microCT imaging postopera...

Watch this Scientific Journal Video about A Surgical Procedure for Resecting the Mouse Rib: A Model for Large-Scale Long Bone Repair at JoVE.com

A Surgical Procedure for Resecting the Mouse Rib: A Model for Large-Scale Long Bone Repair

The overall goal of this procedure is to successfully resect a portion of bone from the rib of a mouse. The procedure was developed as a model to study large-scale long bone repair.

This protocol introduces researchers to a new model for large-scale bone repair utilizing the mouse rib. The procedure details the following: preparation ...

a-surgical-procedure-for-resecting-mouse-rib-model-for-large-scale

Research

JoVE Journal

Medicine

13.0K Views.  Keck School of Medicine, University of Southern California. The overall goal of this procedure is to successfully resect a portion of bone from the rib of a mouse. The procedure was developed as a model to study large-scale long bone repair.

Video: A Surgical Procedure for Resecting the Mouse Rib: A Model for Large-Scale Long Bone Repair

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

Surgical Technique for Spinal Cord Delivery of Therapies: Demonstration of Procedure in Gottingen Minipigs

This is a compact visual description of a combination of surgical technique and device for the delivery of (gene and cell) therapies into the spinal cord. While the technique is demonstrated in the animal, the procedure is FDA-approved and currently being used for stem cell transplantation into the spinal cords of patients with ALS. While the FDA has recognized proof-of-principle data on therapeutic efficacy in highly characterized rodent models, the use of large animals is considered critical for validating the combination of a surgical procedure, a device, and the safety of a final therapy for human use. The size, anatomy, and general vulnerability of the spine and spinal cord of the swine are recognized to better model the human. Moreover, the surgical process of exposing and manipulating the spinal cord as well as closing the wound in the pig is virtually indistinguishable from the human. We believe that the healthy pig model represents a critical first step in the study of procedural safety.

Short visual description of the surgical technique and device used for the delivery of (gene and cell) therapies into the spinal cord. The technique is demonstrated in the animal but is entirely translatable and currently being used for human application.

This is a  compact visual description of a combination of surgical technique and device  for the  delivery of (gene and cell) therapies into the spinal ...

A Mouse Fetal Skin Model of Scarless Wound Repair

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs across species but is age dependent. The transition from a scarless to scarring phenotype occurs in the third trimester of pregnancy in humans and around embryonic day 18 (E18) in mice. However, this varies with the size of the wound with larger defects generating a scar at an earlier gestational age. The emergence of lineage tracing and other genetic tools in the mouse has opened promising new avenues for investigation of fetal scarless wound healing. However, given the inherently high rates of morbidity and premature uterine contraction associated with fetal surgery, investigations of fetal scarless wound healing in vivo require a precise and reproducible surgical model. Here we detail a reliable model of fetal scarless wound healing in the dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

During mammalian development, early gestational skin wounds heal without a scar. Here we detail a reliable and reproducible model of fetal scarless wound healing in the cutaneous dorsum of E16.5 (scarless) and E18.5 (scarring) mouse embryos.

Early in utero, but not in postnatal life, cutaneous wounds undergo regeneration and heal without formation of a scar. Scarless fetal wound healing occurs ...

The Mouse Round-window Approach for Ototoxic Agent Delivery: A Rapid and Reliable Technique for Inducing Cochlear Cell Degeneration

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research involving the cochlea and its associated neural pathways entails exposure of model cochleae to a variety of ototoxic agents. This allows investigators to study the effects of targeted damage to cochlear structures, and in some cases, the self-repair or regeneration of those structures. Various techniques exist for delivery of ototoxic agents to the cochlea. When selecting a particular technique, investigators must consider a number of factors, including the induction of inadvertent systemic toxicity, the amount of cochlear damage produced by the surgical procedure itself, the type of lesion desired, animal survivability, and reproducibility/reliability of results. Currently established techniques include parenteral injection, intra-peritoneal injection, trans-tympanic injection, endolymphatic sac injection, and cochleostomy with perilymphatic perfusion. Each of these methods has been successfully utilized and is well described in the literature; yet, each has various shortcomings. Here, we present a technique for topical application of ototoxic agents directly to the round window niche. This technique is non-invasive to inner ear structures, produces rapid onset of reliably targeted lesions, avoids systemic toxicity, and allows for an intra-animal control (the contra-lateral ear). Results stemming from this approach have helped deeper understanding of auditory pathophysiology, cochlear cell degeneration, and regenerative capacity in response to an acute injury. Future investigations may use this method to conduct interventional studies involving gene therapy and stem cell transplantation to combat hearing loss.

Various methods exist for introducing ototoxic agents to the cochleae of animal models. Presented is a surgical protocol for delivery of ototoxic agents to the round window niche. The procedure is reliable, creates targeted intra-cochlear lesions, and avoids mechanical damage to the microarchitecture. Examination of cochlear self-repair/regeneration is possible.

Investigators have utilized a wide array of animal models and investigative techniques to study the mammalian auditory system. Much of the basic research ...

A Novel Murine Model of Arteriovenous Fistula Failure: The Surgical Procedure in Detail

The arteriovenous fistula (AVF) still suffers from a high number of failures caused by insufficient remodeling and intimal hyperplasia from which the exact pathophysiology remains unknown. In order to unravel the pathophysiology a murine model of AVF-failure was developed in which the configuration of the anastomosis resembles the preferred situation in the clinical setting. A model was described in which an AVF is created by connecting the venous end of the branch of the external jugular vein to the side of the common carotid artery using interrupted sutures. At a histological level, we observed progressive stenotic intimal lesions in the venous outflow tract that is also seen in failed human AVFs. Although this procedure can be technically challenging due to the small dimensions of the animal, we were able to achieve a surgical success rate of 97% after sufficient training. The key advantage of a murine model is the availability of transgenic animals. In view of the different proposed mechanisms that are responsible for AVF failure, disabling genes that might play a role in vascular remodeling can help us to unravel the complex pathophysiology of AVF failure.

Here we present a murine model of arteriovenous fistula (AVF) failure in which a clinically relevant anastomotic configuration is incorporated. This model can be used to study the pathophysiology and to test possible therapeutic interventions.

The arteriovenous fistula (AVF) still suffers from a high number of failures caused by insufficient remodeling and intimal hyperplasia from which the ...

Surgical Angiogenesis in Porcine Tibial Allotransplantation: A New Large Animal Bone Vascularized Composite Allotransplantation Model

Segmental bone loss resulting from trauma, infection malignancy and congenital anomaly remains a major reconstructive challenge. Current therapeutic options have significant risk of failure and substantial morbidity.
Use of bone vascularized composite allotransplantation (VCA) would offer both a close match of resected bone size and shape and the healing and remodeling potential of living bone. At present, life-long drug immunosuppression (IS) is required. Organ toxicity, opportunistic infection and neoplasm risks are of concern to treat such non-lethal indications.
We have previously demonstrated that bone and joint VCA viability may be maintained in rats and rabbits without the need of long-term-immunosuppression by implantation of recipient derived vessels within the VCA. It generates an autogenous, neoangiogenic circulation with measurable flow and active bone remodeling, requiring only 2 weeks of IS. As small animals differ from man substantially in anatomy, bone physiology and immunology, we have developed a porcine bone VCA model to evaluate this technique before clinical application is undertaken. Miniature swine are currently widely used for allotransplantation research, given their immunologic, anatomic, physiologic and size similarities to man. Here, we describe a new porcine orthotopic tibial bone VCA model to test the role of autogenous surgical angiogenesis to maintain VCA viability.
The model reconstructs segmental tibial bone defects using size- and shape-matched allogeneic tibial bone segments, transplanted across a major swine leukocyte antigen (SLA) mismatch in Yucatan miniature swine. Nutrient vessel repair and implantation of recipient derived autogenous vessels into the medullary canal of allogeneic tibial bone segments is performed in combination with simultaneous short-term IS. This permits a neoangiogenic autogenous circulation to develop from the implanted tissue, maintaining flow through the allogeneic nutrient vessels for a short time. Once established, the new autogenous circulation maintains bone viability following cessation of drug therapy and subsequent nutrient vessel thrombosis.

Currently any kind of vascularized composite allotransplantation depends on long-term-immunosuppression, difficult to support for non-life-critical indications. We present a new porcine tibial VCA model that can be used to study bone VCA and demonstrate the use of surgical angiogenesis to maintain bone viability without the need of long-term immune-modulation.

Segmental bone loss resulting from trauma, infection malignancy and congenital anomaly remains a major reconstructive challenge. Current therapeutic ...

Evaluating the Procedure for Performing Awake Cystometry in a Mouse Model

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among laboratories. The goal of this study was to describe the microsurgical procedure used to implant an intravesical tube and the experimental technique for recording urinary bladder pressure in an awake, freely moving mouse. In addition, experimental data is presented to show how surgery, as well as tubing type and size, affect lower urinary tract function and recording sensitivity. The effect of tube diameter on pressure recording was assessed in both polyethylene and polyurethane tubing with different internal diameters. Subsequently, the best performing tube from both materials was surgically implanted into the dome of the urinary bladder of male C57BL/6 mice. Twelve-hour, overnight micturition frequency was recorded in healthy, intact animals and animals 2, 3, 5, and 7 days post-surgery. At harvest, bladders were assessed for signs of swelling using gross observation and were subsequently processed for pathological analysis. The greatest extent of bladder swelling was observed on day 2 and 3, which correlated with behavioral voiding data showing significantly impaired bladder function. By day 5, bladder histology and voiding frequency had normalized. Based on the literature and evidence provided by our studies, we propose the following steps for in vivo recording of intravesical pressure and voided volume in an awake mouse: 1) Perform the surgery using an operating microscope and microsurgical tools, 2) Use polyethylene-10 tubing to minimize movement artifacts, and 3) Perform cystometry on post-operative day 5, when bladder swelling resolves.

This study describes the surgical procedures and experimental techniques for performing awake cystometry in a freely moving mouse. In addition, it provides experimental evidence to support its optimization and standardization.

Awake filling cystometry has been used for a long time to evaluate bladder function in freely moving mice, however, the specific methods used, vary among ...

A Surgical Procedure for the Administration of Drugs to the Inner Ear in a Non-Human Primate Common Marmoset (Callithrix jacchus)

Hearing research has long been facilitated by rodent models, although in some diseases, human symptoms cannot be recapitulated. The common marmoset (Callithrix jacchus) is a small, easy-to-handle New World monkey which has a similar anatomy of the temporal bone, including the middle ear ossicular chains and inner ear to humans, than in comparison with that of rodents. Here, we report a reproducible, safe, and rational surgical approach to the cochlear round window niche for the drug delivery to the inner ear of the common marmoset. We adopted posterior tympanotomy, a procedure used clinically in human surgery, to avoid manipulation of the tympanic membrane that may cause conductive hearing loss. This surgical procedure did not lead to any significant hearing loss. This approach was possible due to the large bulla structure of the common marmoset, although the lateral semicircular canal and vertical portion of the facial nerve should be carefully considered. This surgical method allows us to perform the safe and accurate administration of drugs without hearing loss, which is of great importance in obtaining pre-clinical proof of concept for translational research.

A Surgical Procedure for the Administration of Drugs to the Inner Ear in a Non-Human Primate Common Marmoset Callithrix jacchus

We report a surgical method to administrate drugs to the inner ear of a non-human primate, the common marmoset (Callithrix jacchus), via the round window membrane.

Hearing research has long been facilitated by rodent models, although in some diseases, human symptoms cannot be recapitulated. The common marmoset ...

Standardized Histomorphometric Evaluation of Osteoarthritis in a Surgical Mouse Model

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, primarily in the hip and knee joints, which results in significant impacts on patient mobility and quality of life. To date, there are no existing curative therapies for OA able to slow down or inhibit cartilage degeneration. Presently, there is an extensive body of ongoing research to understand OA pathology and discover novel therapeutic approaches or agents that can efficiently slow down, stop, or even reverse OA. Thus, it is crucial to have a quantitative and reproducible approach to accurately evaluate OA-associated pathological changes in the joint cartilage, synovium, and subchondral bone. Currently, OA severity and progression are primarily assessed using the Osteoarthritis Research Society International (OARSI) or Mankin scoring systems. In spite of the importance of these scoring systems, they are semiquantitative and can be influenced by user subjectivity. More importantly, they fail to accurately evaluate subtle, yet important, changes in the cartilage during the early disease states or early treatment phases. The protocol we describe here uses a computerized and semiautomated histomorphometric software system to establish a standardized, rigorous, and reproducible quantitative methodology for the evaluation of joint changes in OA. This protocol presents a powerful addition to the existing systems and allows for more efficient detection of pathological changes in the joint.

The current protocol establishes a rigorous and reproducible method for quantification of morphological joint changes that accompany osteoarthritis. Application of this protocol can be valuable in monitoring disease progression and evaluating therapeutic interventions in osteoarthritis.

One of the most prevalent joint disorders in the United States, osteoarthritis (OA) is characterized by progressive degeneration of articular cartilage, ...

A Mouse Model for Chronic Pancreatitis via Bile Duct TNBS Infusion

Chronic pancreatitis (CP) is a complex disease involving pancreatic inflammation and fibrosis, glandular atrophy, abdominal pain and other symptoms. Several rodent models have been developed to study CP, of which the bile duct 2,4,6 -trinitrobenzene sulfonic acid (TNBS) infusion model replicates the features of neuropathic pain seen in CP. However, bile duct drug infusion in mice is technically challenging. This protocol demonstrates the procedure of bile duct TNBS infusion for generation of a CP mouse model. TNBS was infused into the pancreas through the ampulla of Vater in the duodenum. This protocol optimized drug volume, surgical techniques, and drug handling during the procedure. TNBS-treated mice showed features of CP as reflected by bodyweight and pancreas weight reductions, changes in pain-associated behaviors, and abnormal pancreatic morphology. With these improvements, mortality associated with TNBS injection was minimal. This procedure is not only critical in generating pancreatic disease models but is also useful in local pancreatic drug delivery.

Chronic pancreatitis (CP) is a disease characterized by inflammation and fibrosis of the pancreas, often associated with intractable abdominal pain. This article focuses on refining the technique to generate a mouse model of CP via bile duct infusion with 2,4,6 -trinitrobenzene sulfonic acid (TNBS).

Chronic pancreatitis (CP) is a complex disease involving pancreatic inflammation and fibrosis, glandular atrophy, abdominal pain and other symptoms. ...