We would like to thank both Mission Connect and the Gillson-Longenbaugh Foundation for providing the funds for this project. We would also like to thank Dr. Catherine Ambrose for her critical advice and use of her DXA equipment. Finally, we would like to thank Dr. Kaori Ono, Department of Integrative Biology and Pharmacology, for her suggestions in DXA femur bone analysis.

1. Mouse spinal transection model
<ol>
 <li>Adult, male, C57BL6 mice (approximately 20-25 g) are anesthetized using a combination of ketamine (200 mg/kg) and xylazine (10 mg/kg). All surgical procedures are performed in an Institutional, IACUC-approved surgical suite under sterile conditions.</li>
 <li>Once deeply anesthetized, the fur of the back is trimmed using electric clippers. The shaved back is first scrubbed with an iodine solution followed by 70% ethanol.</li>
 <li>Prior to the initial incision, the area of the back to be incised is first infiltrated with a local anesthetic (Marcaine) at a concentration of (0.25%, &lt;1ml/kg) to minimize post-surgical pain.</li>
 <li>Using small scissors, a small opening is made in the skin at around the L2 area. This opening is expanded with the same set of scissors, extending longitudinally to the T2 area. The edges of the skin are then held apart via the use of bulldog clamps.</li>
 <li>Micro-scissors are then used to clear muscle tissue from the dorsal spinal lamina at thoracic level 8 (T8). T8 can be identified as described in Kuh and Wrathall (1998) 10. Briefly, T13 can be identified by its dorsal spinous process. Using a pair of #5 Dumont forceps, the last rib can be palpated/identified and counted back to T8. Once cleared, bone Rongeurs are used to perform a dorsal laminectomy at T8.</li>
 <li>Full Spinal Transection Lesion: Once the spinal cord is exposed at T8, an additional set of sterilized micro-scissors are used to sever the spinal cord in the plane perpendicular to the long axis of the cord under a surgical microscope. We use #5 Dumont forceps to gently lift one pole of the spinal cord to confirm the completeness of the lesion.</li>
 <li>Following spinal transection, a small piece of sterile gelfoam, soaked in sterile saline (0.9%) is carefully placed into the lesion cavity to promote hemostasis.</li>
 <li>An additional piece of gelfoam is then placed over the exposed spinal cord. The skin is then closed with sterile, stainless-steel surgical staples. Subjects are then returned to their home cages, placed on a paper towel to prevent aspiration of bedding material and warmed with a heating pad for a period of approximately 12 hours. Subjects are also provided with Hydrogel packs (ClearH2O) and food pellets on the floor of their cages during early recover. Injured subjects are able to access food/water in their home cages once recovered from anesthesia.</li>
 <li>All injured subjects receive twice-daily manual bladder evacuations (at approximately 12 hour intervals) using a modification of the manual method of Crede for the duration of the study (40 days).</li>
 <li>Injured mice are also provided with twice daily intraperitoneal injections of 0.9% saline for three days (0.5 cc) to help maintain hydration, and twice daily injections of the opiate buprenorphine (0.05 mg/kg) to control post-surgical pain for a period of 5 days. If animals exhibit signs of pain after the initial 5 day period, they would receive additional buprenorphine (0.05 mg/kg) daily until signs of pain (reduced mobility, hunched stature, failure to groom, vocalization when handled) have resolved. </li>
</ol>
2. Longitudinal assessment of bone density using the IVIS Lumina XR on the same cohort of spinally-transected mice
<ol>
 <li>Beginning on Day 10 post-SCI and continuing at 10 day intervals until day 40, we assessed right and left femurs and tibiae in living, anesthetized subjects (transection SCI and uninjured age-matched controls).</li>
 <li>On the day of scanning, subjects were transferred in their home cages from the Institution's Vivarium area to the room in which the Caliper IVIS Lumina XR is housed. All subjects are then anesthetized using the same Ketamine/xylazine cocktail previously used in the delivery of SCI. This cocktail assures a state of anesthesia for a period of 1 to 1.5 hours; sufficient for the duration of the scanning procedure.</li>
 <li>The investigator initializes the Lumina XR device and allows the internal camera to reach operational temperature (-90&deg;C) (~10 minutes).</li>
 <li>Once anesthetized, the subject (transection or control) is gently placed on the animal platform within the Lumina XR. A high magnification lens is inserted into the device to allow focus on both femur and tibial regions (Field of view 2.4X2.4cm with high-magnification lens). If the subject is improperly positioned outside of the field of view, the door is opened and the subject re-positioned until the left or right hind limb is centered (See Figure 2a for proper placement of the hind limb).</li>
 <li>Once properly positioned, the X-ray function can be executed. Make a selection from the Energy drop-down list suitable for Animal (Living subject, 35 Kv 100uA, filtered X-Rays)</li>
 <li>Once the animal is in the right position, enable the X-ray feature by check-marking X-Ray in the control panel. Acquire the X-ray image. Make sure that the entire femur and tibia are visible (see Figure 2b). Raw image data is automatically saved to the hard drive. Representative .TIFF files are also saved. The mouse is then returned to its home cage and allowed to recover under investigator observation. The process is then repeated with the next mouse. 
 Note: The Living Image software displays transformed X-ray images by default. To display raw X-ray images, remove the check mark next to X-Ray Absorption in the Corrections/Filtering tools. When the X-ray data has been corrected for absorption, you can evaluate relative bone density by comparing the signal intensities of measurement ROIs. The ROI intensity increases with increasing tissue density.</li>
</ol>
3. Image analysis of IVIS X-ray scans
<ol>
 <li>Open the software by double-clicking on the Living Image Icon.</li>
 <li>Load an X- ray image by clicking the Browse button. In the dialog box that appears, select the folder of interest and click OK. The Living Image browser displays the selected data along with the user ID, label information, and camera configuration information. To open data, do one of the following: Double-click the data row, Right-click the data name and select Load on the shortcut menu, Select the data row and click Load, or Double-click the thumbnail. The image and Tool Palette are displayed. Open data is highlighted in green in the browser.</li>
 <li>Click on ROI Tools in the Tool Palette. In the ROI Tools, select Measurement ROI from the Type drop-down list. To load the 3 ROI's used in this experiment, click the Square icon and load 3 squares.</li>
 <li>Using a ruler, measure the length of the femur. Adjust the length of two of the squares to be 1/8th of the total femur length. Adjust the width of these two squares to be 1/24th the total femur length. With your ruler, measure 1/8th of the distance from the proximal end of the femur, and situate the square so that it is centered within the femur. Situate the second square so that it lies 1/4th of the total femur length from the distal end of the femur (Fig. 3). These ROI's can be used to measure both proximal and distal femur regions.</li>
 <li>Using the ruler, measure the length of the tibia. Adjust the length of the third square to be 1/8th of the total tibial length. Adjust the width to 1/30th of the total tibial length. Situate the square so that is centered and 1/8th the total length of the tibia distance from the proximal end of the tibia (see Figure 3).</li>
 <li>Click on the Measure icon (a pencil and ruler). The ROI intensity measurements appear in the X-ray image and the ROI measurements table appears. Export this table to your desired location as a .csv file. This will allow you to open the table using excel.</li>
 <li>Repeat this with all of your saved images.</li>
 <li>Consolidate all of your data onto one excel sheet. Statistical significance was determined via student's t-test using either Microsoft Excel or SigmaPlot 11.0 software (Systat Software).</li>
</ol>
4. Post-mortem analysis of bone density:
<ol>
 <li>Following the acquisition of the final longitudinal X-ray scan within the IVIS Lumina XR, mice are subsequently deeply anesthetized with Beuthanasia (75 mg pentobarbital/kg). Once deep anesthesia has been achieved, the mice are transcardially perfused with iced-cold phosphate-buffered saline with heparain (40 mg/liter) to exsanguinate.</li>
 <li>Once exsanguinated, excise both femurs. Pay special care to remove as much soft tissue as possible, for this has been shown to affect density measurements 11. Wrap the femurs with water soaked gauze and store at -20C until you are ready to analyze them.</li>
</ol>
5. DXA Analysis using a Hologic QDR 4000 Bone Densitometer
<ol>
 <li>Thaw gauze-soaked femurs and transfer to Bone Densitometer suite.</li>
 <li>Calibrate the apparatus according to the manufacturer's protocol; ensure that the BMC and BMD values fall within the acceptable limits.</li>
 <li>Place a brass collimator in the machine. This allows the user to limit the size and angle of the X-ray beam to focus on a specific target.</li>
 <li>Submerge the thawed femur into a Petri dish filled with water (condyles to the left, with the femur parallel to the bed), and position it just to the right of the laser.</li>
 <li>Enter a Biography (a descriptive title that includes animal identification information, treatment, etc.) for the femur you are about to scan.</li>
 <li>Enter Scan from the Selections Menu, then select regionA hi-res. Indicate your scanning parameters: a scanning region of 2 x 0.7489 inches, with 0.01 inch line spacing and 0.00499 inch point resolution. 
 Note: Monitor the scan while it is in progress. As the X-ray beam raster scans the sample, monitor to ensure that there is sufficient water in the chamber to completely cover the bone sample.</li>
 <li>Enter Analyze under the Analysis Selection menu. Follow the prompts to highlight the entire femur as an ROI. A report page will come up with the calculated BMC (grams) and BMD (gms/cm2).</li>
 <li>Repeat these steps to analyze the remaining femurs.</li>
 <li>Consolidate all of your data onto one excel sheet and perform statistical analysis (t-test).</li>
</ol>
6. Representative Results:
The relative bone density loss of a mouse tibia and femur after spinal cord injury when compared to na&iuml;ve mice is detectable using the above method. There is a detectable significant decrease in bone density after just 10 days (12%, p&lt;0.0005), with up to 15% bone density loss at 40 days (p&lt;0.0005, Figure 4). Bone density loss in the femur was observed at 40 days post injury (7% decrease, p&lt;0.05, Figure 5). These results provide evidence for the use of non-invasive x-ray imaging for the longitudinal observation of bone density change after spinal cord injury.
In order to compare the efficacy of this method to what is currently available; we analyzed the excised femurs of these mice 40 days after injury using DXA imaging. A representation of the data output can be seen in Figure 6. We found that there was a significant loss of bone mineral content in the SCI mice when compared to na&iuml;ve (12% decrease, p&lt;0.05, Figure 7). Bone mineral density did not change significantly, but did follow a similar trend (Figure 8). These results are similar to those found in the literature; Picard et al observed a 13.5% decrease (p&lt;0.001) in BMC, but no significant decrease in BMD (Picard 2008).
<img alt="Figure 1" src="/files/ftp_upload/3246/3246fig1.jpg" /> 
Figure 1. Experimental Timeline.
<img alt="Figure 2" src="/files/ftp_upload/3246/3246fig2.jpg" /> 
Figure 2. Representative orientation of left hind-limb: a) photograph and b) x-ray.
<img alt="Figure 3" src="/files/ftp_upload/3246/3246fig3.jpg" /> 
Figure 3. ROI dimensions and orientation within the regions of the proximal femur and tibia.
<img alt="Figure 4" src="/files/ftp_upload/3246/3246fig4.jpg" /> 
Figure 4. Bone density loss after SCI in the proximal tibia 10, 20, 30 and 40 days after injury (n=5) compared to age-matched naives (n=5). Error bars represent SEM; **p&lt;0.005, *** p&lt;0.0005.
<img alt="Figure 5" src="/files/ftp_upload/3246/3246fig5.jpg" /> 
Figure 5. Bone density loss after SCI in the proximal femur 10, 20, 30 and 40 days after injury (n=5) compared to age-matched naives (n=5). Error bars represent SEM; *p&lt;0.05.
<img alt="Figure 6" src="/files/ftp_upload/3246/3246fig6.jpg" /> 
Figure 6. Bone density loss after SCI in the distal femur 10, 20, 30 and 40 days after injury (n=5) compared to age-matched na&iuml;ve controls (n=5). Error bars represent SEM; *p&lt;0.01-0.05; **p&lt;0.001-0.01; ***p&lt;0.0001-0.001; ****p&lt;0.0001.
<img alt="Figure 7" src="/files/ftp_upload/3246/3246fig7.jpg" /> 
Figure 7. Representative image of DXA data showing BMC and BMD output.
<img alt="Figure 8" src="/files/ftp_upload/3246/3246fig8.jpg" /> 
Figure 8. DXA analysis of bone mineral content (grams) in the femurs of SCI mice 40 days post injury (n=5) vs. age-matched naives (n=5). Error bars represent SEM; *p&lt;0.05.
<img alt="Figure 9" src="/files/ftp_upload/3246/3246fig9.jpg" /> 
Figure 9. DXA analysis of bone mineral density (mg/cm2) in the femurs of SCI mice 40 days post injury (n=5) vs. age-matched naives (n=5). Error bars represent SEM; no significant difference.

<ol>
	<li>Jiang, S. D., Jiang, L. S., Dai, L. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+osteoporosis+in+spinal+cord+injury.">Mechanisms of osteoporosis in spinal cord injury.</a> Clin. Endocrinol. (Oxford). 65, 555-565 (2006).</li><li>Uebelhart, D., Demiaux-Domenech, B., Roth, M., Chantraine, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bone+metabolism+in+spinal+cord+injured+individuals+and+in+others+who+have+prolonged+immobilisation.+A+review.">Bone metabolism in spinal cord injured individuals and in others who have prolonged immobilisation. A review.</a> Paraplegia. 33, 669-673 (1995).</li><li>Elefteriou, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+signaling+and+the+regulation+of+bone+remodeling.">Neuronal signaling and the regulation of bone remodeling.</a> Cell. Mol. Life. Sci. 62, 2339-2349 (2005).</li><li>Finsen, V., Indredavik, B., Fougner, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bone+mineral+and+hormone+status+in+paraplegics.">Bone mineral and hormone status in paraplegics.</a> Paraplegia. 30, 343-347 (1992).</li><li>Sugawara, H., Linsenmeyer, T. A., Beam, H., Parsons, 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=Mechanical+properties+of+bone+in+a+paraplegic+rat+model.">Mechanical properties of bone in a paraplegic rat model.</a> J. Spinal. Cord. Med. 21, 302-308 (1998).</li><li>Kiratli, B. J., Smith, A. E., Nauenberg, T., Kallfelz, C. F., Perkash, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bone+mineral+and+geometric+changes+through+the+femur+with+immobilization+due+to+spinal+cord+injury.">Bone mineral and geometric changes through the femur with immobilization due to spinal cord injury.</a> J. Rehabil. Res. Dev. 37, 225-233 (2000).</li><li>Shen, J., Fan, L., Yang, J., Shen, A. G., Hu, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+longitudinal+Raman+microspectroscopic+study+of+osteoporosis+induced+by+spinal+cord+injury.">A longitudinal Raman microspectroscopic study of osteoporosis induced by spinal cord injury.</a> Osteoporos. Int. 21, 81-87 (2010).</li><li>Guertin, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technological+platform+to+optimize+combinatorial+treatment+design+and+discovery+for+chronic+spinal+cord+injury.">A technological platform to optimize combinatorial treatment design and discovery for chronic spinal cord injury.</a> J. Neurosci. Res. 86, 3039-3051 (2008).</li><li>Picard, S., Lapointe, N. P., Brown, J. P., Guertin, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Histomorphometric+and+densitometric+changes+in+the+femora+of+spinal+cord+transected+mice.">Histomorphometric and densitometric changes in the femora of spinal cord transected mice.</a> Anat. Rec. (Hoboken). 291, 303-307 (2008).</li><li>Kuhn, P. L., Wrathall, 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=A+mouse+model+of+graded+contusive+spinal+cord+injury.">A mouse model of graded contusive spinal cord injury.</a> J. Neurotrauma. 15, 125-140 (1998).</li><li>Tsujio, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=mineral+analysis+through+dual+energy+X-ray+absorptiometry+in+laboratory+animals.">mineral analysis through dual energy X-ray absorptiometry in laboratory animals.</a> J. Vet. Med. Sci. 71, 1493-1497 (2009).</li></ol>

Free Access to this article is sponsored by Caliper Life Sciences.

This study presents a novel, non-invasive method for assessing density changes within individual bones (tibiae and femurs) longitudinally, within a single cohort of mice, using the Caliper IVIS Lumina XR imaging system. SCI produced a decrease in bone density in both femurs and tibiae when studied within a single cohort of spinally-transected mice over a 40 day assay period compared to age-matched, uninjured, control mice. Our results in the femur were confirmed, post-mortem, through the use of DXA, suggesting that the application of the Caliper Instruments IVIS Lumina XR can serve as a sensitive measuring tool to assess the long-term and progressive effects of SCI on bone loss.
One potential benefit to investigators using this system is the ability to longitudinally follow the development of pathological changes in the hind limb bones of animals following SCI. The ability to assess a single cohort over long time periods provides strong advantages to the investigator in terms of: 1) costs associated with animal purchase and long-term care. Instead of a study requiring large numbers of animals that need to be sacrificed at specific time points in order to assess time-dependent changes, these pathological outcomes can be followed within the same cohort over that same time period, 2) variability: in addition to comparison with control cohorts, the outcomes generated within the single cohort of injured subjects can be assessed and compared between time points for the same animals throughout the extent of the study; once again, reducing both intra-animal variability as well as the need for larger cohorts of experimental subjects.
While the Caliper IVIS Lumina XR provided us with important longitudinal information regarding the development of SCI-induced osteoporosis, there are limitations that should be considered: 1) while this platform has allowed us to generate high-resolution X-ray data in the hind limb bones of mice following SCI, its utility may be limited to small rodents such as mice. The maximum subject height allowed within the Lumina XR is 2.8 cm. Anything above that height cannot be successfully imaged using the X-ray component of the Lumina XR. In our hands, this has precluded our ability to study rats in the 200-250 gram weight. While it may be possible to image rats under this weight range or other rodents such as hamsters or gerbils, this will require additional testing by the PI. 2) Joint orientation. It is critical that the Investigator carefully arrange the limbs in an easily reproducible orientation that will allow successful comparison between imaging sessions. Standardization of limb placement will allow the Investigator to generate and apply standardized ROI's that can be saved and applied across both subjects and time.
Overall, we believe that the IVIS Lumina XR provides an excellent platform with which to model the development of SCI-dependent osteoporosis in mice. The ability to study bone loss longitudinally within the same mouse cohort will allow us to: 1) better understand the temporal nature of bone loss and demineralization after SCI, 2) determine whether these changes stabilize over time, 3) explore, with the availability of both transgenic and null mouse lines, the ability to study the molecular mechanisms underlying these pathological changes in a reductionist manner, and 4) quite possibly the most important advantage, begin to test novel interventions designed to either prevent the development of osteoporosis in the early stages of injury, or, to reverse such changes once osteoporosis has already developed. Finally, the Lumina XR, in addition to providing excellent X-ray imaging potential, can also be used to study both bioluminescent and fluorescent signals in living animals. One could easily imagine incorporating the other modalities offered within this system to look at a multitude of factors that determine bone dynamics mechanistically following SCI or any other disease process that results in osteoporosis.

<table border="1">
<tbody>
 <tr>
 <th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 <th>Comments</th>
 </tr>
 <tr>
 <td>Beuthanasia</td>
 <td>Butler Schein Animal Health</td>
 <td>001848</td>
 <td></td>
 </tr>
 <tr>
 <td>Ketathesia (Ketamine Hydrochloride)</td>
 <td>Butler Schein Animal Health</td>
 <td>023061</td>
 <td></td>
 </tr>
 <tr>
 <td>Xylazine</td>
 <td>Butler Schein Animal Health</td>
 <td>037849</td>
 <td></td>
 </tr>
 <tr>
 <td>Bupivicaine</td>
 <td>Butler Schein Animal Health</td>
 <td>021801</td>
 <td></td>
 </tr>
 <tr>
 <td>Gelfoam; 7MM</td>
 <td>Fisher Scientific</td>
 <td>NC0085178</td>
 <td>To promote hemostasis during surgery</td>
 </tr>
 <tr>
 <td><a href="http://www.caliperls.com/products/preclinical-imaging/ivis-lumina-xr.htm">IVIS Lumina XR</a></td>
 <td>Caliper Life Sciences</td>
 <td>135400</td>
 <td></td>
 </tr>
 <tr>
 <td><a href="http://www.caliperls.com/products/preclinical-imaging/ivis-lumina-xr.htm">ZFOV</a>, 2.5 cm field of view lens</td>
 <td>Caliper Life Sciences</td>
 <td>127285</td>
 <td>Absolutely necessary to generate high magnification X-ray images of bone structure</td>
 </tr>
 <tr>
 <td>QDR Bone Densitometer</td>
 <td>Hologic</td>
 <td></td>
 <td>Model used no longer in production</td>
 </tr>
</tbody>
</table>

longitudinal evaluation of mouse hind limb bone loss after spinal cord injury using novel, in vivo, methodology

Spinal cord injury (SCI) is often accompanied by osteoporosis in the sublesional regions of the pelvis and lower extremities, leading to a higher frequency of fractures 1. As these fractures often occur in regions that have lost normal sensory function, the patient is at a greater risk of fracture-dependent pathologies, including death. SCI-dependent loss in both bone mineral density (BMD, grams/cm2) and bone mineral content (BMC, grams) has been attributed to mechanical disuse 2, aberrant neuronal signaling 3 and hormonal changes 4. The use of rodent models of SCI-induced osteoporosis can provide invaluable information regarding the mechanisms underlying the development of osteoporosis following SCI as well as a test environment for the generation of new therapies 5-7 (and reviewed in 8). Mouse models of SCI are of great interest as they permit a reductionist approach to mechanism-based assessment through the use of null and transgenic mice. While such models have provided important data, there is still a need for minimally-invasive, reliable, reproducible, and quantifiable methods in determining the extent of bone loss following SCI, particularly over time and within the same cohort of experimental animals, to improve diagnosis, treatment methods, and/or prevention of SCI-induced osteoporosis.
An ideal method for measuring bone density in rodents would allow multiple, sequential (over time) exposures to low-levels of X-ray radiation. This study describes the use of a new whole-animal scanner, the IVIS Lumina XR (Caliper Instruments) that can be used to provide low-energy (1-3 milligray (mGy)) high-resolution, high-magnification X-ray images of mouse hind limb bones over time following SCI. Significant bone density loss was seen in the tibiae of mice by 10 days post-spinal transection when compared to uninjured, age-matched control (na&iuml;ve) mice (13% decrease, p&lt;0.0005). Loss of bone density in the distal femur was also detectable by day 10 post-SCI, while a loss of density in the proximal femur was not detectable until 40 days post injury (7% decrease, p&lt;0.05). SCI-dependent loss of mouse femur density was confirmed post-mortem through the use of Dual-energy X-ray Absorptiometry (DXA), the current "gold standard" for bone density measurements. We detect a 12% loss of BMC in the femurs of mice at 40 days post-SCI using the IVIS Lumina XR. This compares favorably with a previously reported BMC loss of 13.5% by Picard and colleagues who used DXA analysis on mouse femurs post-mortem 30 days post-SCI 9. Our results suggest that the IVIS Lumina XR provides a novel, high-resolution/high-magnification method for performing long-term, longitudinal measurements of hind limb bone density in the mouse following SCI.

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Longitudinal Evaluation of Mouse Hind Limb Bone Loss After Spinal Cord Injury using Novel, in vivo, Methodology

A longitudinal examination of bone loss in the femurs and tibiae of adult mice was performed following spinal cord injury using sequential low-dose X-ray scans. Tibia bone loss was detected throughout the study, while bone loss in the femur was not detected until 40 days post injury.

Longitudinal Evaluation of Mouse Hind Limb Bone Loss After Spinal Cord Injury using Novel, in vivo, Methodology

Spinal cord injury (SCI) is often accompanied by osteoporosis in the sublesional regions of the pelvis and lower extremities, leading to a higher ...

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15.1K 次观看.  University of Texas Health Science Center at Houston. 在成年小鼠股骨和胫骨的纵向检查的骨质流失，脊髓损伤后使用连续低剂量X射线扫描。整个研究过程中胫骨损耗检测，直到检测到伤后40天，而在股骨的骨损失。

视频: Longitudinal Evaluation of Mouse Hind Limb Bone Loss After Spinal Cord Injury using Novel, in vivo, Methodology

Acute and Chronic Tactile Sensory Testing after Spinal Cord Injury in Rats

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory testing in rat SCI models is needed. However, until recently, no single testing approach had been validated for SCI so that standardized methods have not been implemented across labs. Additionally, available testing methods could not be implemented acutely or when severe motor impairments existed, preventing studies of the development of SCI-induced allodynia3. Here we present two validated sensory testing methods using von Frey Hair (VFH) monofilaments which quantify changes in tactile sensory thresholds after SCI4-5. One test is the well-established Up-Down test which demonstrates high sensitivity and specificity across different SCI severities when tested chronically5. The other test is a newly-developed dorsal VFH test that can be applied acutely after SCI when allodynia develops, prior to motor recovery4-5. Each VFH monofilament applies a calibrated force when touched to the skin of the hind paw until it bends. In the up-down method, alternating VFHs of higher or lower forces are used on the plantar L5 dermatome to delineate flexor withdrawal thresholds. Successively higher forces are applied until withdrawal occurs then lower force VFHs are used until withdrawal ceases. The tactile threshold reflects the force required to elicit withdrawal in 50% of the stimuli. For the new test, each VFH is applied to the dorsal L5 dermatome of the paw while the rat is supported by the examiner. The VFH stimulation occurs in ascending order of force until at least 2 of 3 applications at a given force produces paw withdrawal. Tactile sensory threshold is the lowest force to elicit withdrawal 66% of the time. Acclimation, testing and scoring procedures are described. Aberrant trials that require a retest and typical trials are defined. Animal use was approved by Ohio State University Animal Care and Use Committee.

We describe two tactile sensory testing methods for acute or chronic periods of spinal cord injury in rats. These validated procedures can detect the development and maintenance of allodynia-like sensations.

Spinal cord injury (SCI) impairs sensory systems causing allodynia1-8. To identify cellular and molecular causes of allodynia, sensitive and valid sensory ...

Controlled Cervical Laceration Injury in Mice

Use of genetically modified mice enhances our understanding of molecular mechanisms underlying several neurological disorders such as a spinal cord injury (SCI). Freehand manual control used to produce a laceration model of SCI creates inconsistent injuries often associated with a crush or contusion component and, therefore, a novel technique was developed. Our model of cervical laceration SCI has resolved inherent difficulties with the freehand method by incorporating 1) cervical vertebral stabilization by vertebral facet fixation, 2) enhanced spinal cord exposure, and 3) creation of a reproducible laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm in depth without associated contusion. Compared to the standard methods of creating a SCI laceration such as freehand use of a scalpel or scissors, our method has produced a consistent lesion. This method is useful for studies on axonal regeneration of corticospinal, rubrospinal, and dorsal ascending tracts.

A novel technique to create a reproducible in vivo model of cervical spinal cord laceration injury in the mouse is described. This technique is based on spine stabilization by fixation of the cervical facets and laceration of the spinal cord using an oscillating blade with an accuracy of &plusmn;0.01 mm.

Use of genetically modified mice enhances our  understanding of molecular mechanisms underlying several neurological disorders  such as a spinal cord ...

A Contusion Model of Severe Spinal Cord Injury in Rats

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is described to obtain a consistent model of severe SCI. Use of a stereotactic frame and computer controlled impactor allows for creation of reproducible injury. Hypothermia and urinary tract infection pose significant challenges in the post-operative period. Careful monitoring of animals with daily weight recording and bladder expression allows for early detection of post-operative complications. The functional results of this contusion model are equivalent to transection models. The contusion model can be utilized to evaluate the efficacy of both neuroprotective and neuroregenerative approaches.

A contusion model of severe spinal cord injury is described. Detailed pre-operative, operative and post-operative steps are described to obtain a consistent model.

The translational potential of novel treatments should be investigated in severe spinal cord injury (SCI) contusion models. A detailed methodology is ...

A Novel Vertebral Stabilization Method for Producing  Contusive Spinal Cord Injury

Clinically-relevant animal cervical spinal cord injury (SCI) models are essential for developing and testing potential therapies; however, producing reliable cervical SCI is difficult due to lack of satisfactory methods of vertebral stabilization. The conventional method to stabilize the spine is to suspend the rostral and caudal cervical spine via clamps attached to cervical spinous processes. &#160;However, this method of stabilization fails to prevent tissue yielding during the contusion as the cervical spinal processes are too short to be effectively secured by the clamps (Figure 1). &#160;Here we introduce a new method to completely stabilize the cervical vertebra at the same level of the impact injury. &#160;This method effectively minimizes movement of the spinal column at the site of impact, which greatly improves the production of consistent SCIs. &#160;We provide visual description of the equipment (Figure 2-4), methods, and a step-by-step protocol for the stabilization of the cervical 5 vertebra (C5) of adult rats, to perform laminectomy (Figure 5) and produce a contusive SCI thereafter. &#160;Although we only demonstrate a cervical hemi-contusion using the NYU/MASCIS impactor device, this vertebral stabilization technique can be applied to other regions of the spinal cord, or be adapted to other SCI devices. &#160;Improving spinal cord exposure and fixation through vertebral stabilization may be valuable for producing consistent and reliable injuries to the spinal cord. &#160;This vertebral stabilization method can also be used for stereotactic injections of cells and tracers, and for imaging using two-photon microscopy in various neurobiological studies.

Vertebral stabilization is necessary for minimizing variability, and for producing consistent experimental spinal cord injuries.&#160; Using a customized stabilizing apparatus in conjunction with the NYU/MASCIS impactor device, we have demonstrated here the proper equipment and procedure for generating reproducible hemi-contusive cervical (C5) spinal cord injuries in adult rats.

Clinically-relevant animal cervical spinal cord injury (SCI) models are essential for developing and testing potential therapies; however, producing ...

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment (RMCA) in Individuals with Chronic Spinal Cord Injury

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination is disrupted by spinal cord injury (SCI), control of respiratory muscles innervated below the injury level is compromised1,2 leading to respiratory muscle dysfunction and pulmonary complications. These conditions are among the leading causes of death in patients with SCI3. Standard pulmonary function tests that assess respiratory motor function include spirometrical and maximum airway pressure outcomes: Forced Vital Capacity (FVC), Forced Expiratory Volume in one second (FEV1), Maximal Inspiratory Pressure (PImax) and Maximal Expiratory Pressure (PEmax)4,5. These values provide indirect measurements of respiratory muscle performance6. In clinical practice and research, a surface electromyography (sEMG) recorded from respiratory muscles can be used to assess respiratory motor function and help to diagnose neuromuscular pathology. However, variability in the sEMG amplitude inhibits efforts to develop objective and direct measures of respiratory motor function6. Based on a multi-muscle sEMG approach to characterize motor control of limb muscles7, known as the voluntary response index (VRI)8, we developed an analytical tool to characterize respiratory motor control directly from sEMG data recorded from multiple respiratory muscles during the voluntary respiratory tasks. We have termed this the Respiratory Motor Control Assessment (RMCA)9. This vector analysis method quantifies the amount and distribution of activity across muscles and presents it in the form of an index that relates the degree to which sEMG output within a test-subject resembles that from a group of healthy (non-injured) controls. The resulting index value has been shown to have high face validity, sensitivity and specificity9-11. We showed previously9 that the RMCA outcomes significantly correlate with levels of SCI and pulmonary function measures. We are presenting here the method to quantitatively compare post-spinal cord injury respiratory multi-muscle activation patterns to those of healthy individuals.

Evaluation of Respiratory Muscle Activation Using Respiratory Motor Control Assessment RMCA in Individuals with Chronic Spinal Cord Injury

The purpose of this publication is to present our original work on a multi-muscle surface electromyographic approach to quantitatively characterize respiratory muscle activation patterns in individuals with chronic spinal cord injury using vector-based analysis.

During breathing, activation of respiratory muscles is coordinated by integrated input from the brain, brainstem, and spinal cord. When this coordination ...

In vivo Macrophage Imaging Using MR Targeted Contrast Agent for Longitudinal Evaluation of Septic Arthritis

Macrophages are key-cells in the initiation, the development and the regulation of the inflammatory response to bacterial infection. Macrophages are intensively and increasingly recruited in septic joints from the early phases of infection and the infiltration is supposed to regress once efficient removal of the pathogens is obtained. The ability to identify in vivo macrophage activity in an infected joint can therefore provide two main applications: early detection of acute synovitis and monitoring of therapy.
In vivo noninvasive detection of macrophages can be performed with magnetic resonance imaging using iron nanoparticles such as ultrasmall superparamagnetic iron oxide (USPIO). After intravascular or intraarticular administration, USPIO are specifically phagocytized by activated macrophages, and, due to their magnetic properties, induce signal changes in tissues presenting macrophage infiltration. A quantitative evaluation of the infiltrate is feasible, as the area with signal loss (number of dark pixels) observed on gradient echo MR images after particles injection is correlated with the amount of iron within the tissue and therefore reflects the number of USPIO-loaded cells.
We present here a protocol to perform macrophage imaging using USPIO-enhanced MR imaging in an animal model of septic arthritis, allowing an initial and longitudinal in vivo noninvasive evaluation of macrophages infiltration and an assessment of therapy action.

In vivo Macrophage Imaging Using MR Targeted Contrast Agent for Longitudinal Evaluation of Septic Arthritis

We demonstrate how to perform macrophage MR imaging using ultrasmall superparamagnetic contrast agent (USPIO) in septic arthritis, allowing an initial and longitudinal in vivo non-invasive evaluation of macrophages infiltration and an assessment of therapy efficacy.

Macrophages are key-cells in the initiation, the development and the regulation of the inflammatory response to bacterial infection. Macrophages are ...

Controlled Cortical Impact Model for Traumatic Brain Injury

Every year over a million Americans suffer a traumatic brain injury (TBI). Combined with the incidence of TBIs worldwide, the physical, emotional, social, and economical effects are staggering. Therefore, further research into the effects of TBI and effective treatments is necessary. The controlled cortical impact (CCI) model induces traumatic brain injuries ranging from mild to severe. This method uses a rigid impactor to deliver mechanical energy to an intact dura exposed following a craniectomy. Impact is made under precise parameters at a set velocity to achieve a pre-determined deformation depth. Although other TBI models, such as weight drop and fluid percussion, exist, CCI is more accurate, easier to control, and most importantly, produces traumatic brain injuries similar to those seen in humans. However, no TBI model is currently able to reproduce pathological changes identical to those seen in human patients. The CCI model allows investigation into the short-term and long-term effects of TBI, such as neuronal death, memory deficits, and cerebral edema, as well as potential therapeutic treatments for TBI.

Traumatic brain injuries (TBIs) remain a serious health problem. Using the controlled cortical impact surgery model, research on the effects of TBI and possible treatment methods may be performed.

Every year over a million Americans suffer a traumatic brain injury (TBI).  Combined with the incidence of TBIs worldwide, the physical, emotional, ...

A Radio-telemetric System to Monitor Cardiovascular Function in Rats with Spinal Cord Transection and Embryonic Neural Stem Cell Grafts

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter connected to a radio transmitter into the femoral artery of rats that underwent a T4 spinal cord transection with or without grafting of embryonic brainstem-derived neural stem cells expressing green fluorescent protein. Compared to other methods such as cannula insertion or tail-cuff, telemetry is advantageous to continuously monitor blood pressure and heart rate in freely moving animals. It is also capable of long term multiple data acquisitions. In spinal cord injured rats, basal cardiovascular data under unrestrained condition and autonomic dysreflexia in response to colorectal distension were successfully recorded. In addition, cardiovascular parameters before and after SCI can be compared in the same rat if a transmitter is implanted before a spinal cord transection. One limitation of the described telemetry procedure is that implantation in the femoral artery may influence the blood supply to the ipsilateral hindlimb.

We present a protocol for using a radio-telemetric system to record cardiovascular parameters in T4 spinal cord transected rats eight weeks after embryonic brainstem neural stem cell grafting into the lesion site. Telemetry is an advanced technique to accurately evaluate cardiovascular function in conscious freely moving spinal cord injured rats.

High thoracic or cervical spinal cord injury (SCI) can lead to cardiovascular dysfunction. To monitor cardiovascular parameters, we implanted a catheter ...

Calibrated Forceps Model of Spinal Cord Compression Injury

Compression injuries of the murine spinal cord are valuable animal models for the study of spinal cord injury (SCI) and spinal regenerative therapy. The calibrated forceps model of compression injury is a convenient, low cost, and very reproducible animal model for SCI. We used a pair of modified forceps in accordance with the method published by Plemel et al. (2008) to laterally compress the spinal cord to a distance of 0.35 mm. In this video, we will demonstrate a dorsal laminectomy to expose the spinal cord, followed by compression of the spinal cord with the modified forceps. In the video, we will also address issues related to the care of paraplegic laboratory animals. This injury model produces mice that exhibit impairment in sensation, as well as impaired hindlimb locomotor function. Furthermore, this method of injury produces consistent aberrations in the pathology of the SCI, as determined by immunohistochemical methods. After watching this video, viewers should be able to determine the necessary supplies and methods for producing SCI of various severities in the mouse for studies on SCI and/or treatments designed to mitigate impairment after injury.

Spinal cord injury models should be highly reproducible. We demonstrate that the calibrated forceps compression model of spinal cord injury is an easy to use surgical method for generating reproducible injuries to the murine spinal cord.

Compression injuries of the murine spinal cord are valuable animal models for the study of spinal cord injury (SCI) and spinal regenerative therapy. The ...

Contrast Enhanced Ultrasound Imaging for Assessment of Spinal Cord Blood Flow in Experimental Spinal Cord Injury

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an important target for neuroprotective therapies. Although several techniques have been described to assess SCBF, they all have significant limitations. To overcome the latter, we propose the use of real-time contrast enhanced ultrasound imaging (CEU). Here we describe the application of this technique in a rat contusion model of SCI. A jugular catheter is first implanted for the repeated injection of contrast agent, a sodium chloride solution of sulphur hexafluoride encapsulated microbubbles. The spine is then stabilized with a custom-made 3D-frame and the spinal cord dura mater is exposed by a laminectomy at ThIX-ThXII. The ultrasound probe is then positioned at the posterior aspect of the dura mater (coated with ultrasound gel). To assess baseline SCBF, a single intravenous injection (400 &#181;l) of contrast agent is applied to record its passage through the intact spinal cord microvasculature. A weight-drop device is subsequently used to generate a reproducible experimental contusion model of SCI. Contrast agent is re-injected 15 min following the injury to assess post-SCI SCBF changes. CEU allows for real time and in-vivo assessment of SCBF changes following SCI. In the uninjured animal, ultrasound imaging showed uneven blood flow along the intact spinal cord. Furthermore, 15 min post-SCI, there was critical ischemia at the level of the epicenter while SCBF remained preserved in the more remote intact areas. In the regions adjacent to the epicenter (both rostral and caudal), SCBF was significantly reduced. This corresponds to the previously described &#8220;ischemic penumbra zone&#8221;. This tool is of major interest for assessing the effects of therapies aimed at limiting ischemia and the resulting tissue necrosis subsequent to SCI.

Contrast Enhanced Ultrasound imaging is a reliable in-vivo tool for quantifying spinal cord blood flow in an experimental rat spinal cord injury model. This paper contains a comprehensive protocol for application of this technique in association with a contusion model of thoracic spinal cord injury.

Reduced spinal cord blood flow (SCBF) (i.e., ischemia) plays a key role in traumatic spinal cord injury (SCI) pathophysiology and is accordingly an ...