This work was supported by the National Natural Science Foundation of China (81973607) and Essential Drug Research and Development (2019ZX09201004-003-032) from Ministry of Science and Technology of China.

The investigations described conform to the Guidelines for the Care and Use of Laboratory Animals of the National Institutes of Health and were approved by Shanghai University of Traditional Chinese Medicine Animal Care and Use Committee. All surgical manipulations were performed under deep anesthesia and the animals did not experience pain at any stage during the procedure.
1. Pre-operation preparation
<ol>
	<li>Instrument sterilization: Steam-sterilize surgical instruments in an autoclave (121 &#176;C for 15 min) prior to the surgery. Pack instruments in a metal container and maintain them until they are used in the surgery.</li>
	<li>Surgery platform setup: Assign a bench area of at least 60 cm x 60 cm for the operation. Clean the surface of area with 75% alcohol and cover with a disposable medical towel. Place a sterile surgical instruments pack, reagents, surgical items onto a disposable medical towel within the upper 1/3 of area. Leave the remaining 2/3 of area clean for surgical operation.&#160;Add a hotpad underneath surgical pad for thermal support.</li>
	<li>Animal preparation
	<ol>
		<li>Place the animal (C57BL/6J mice, male, 8-week old) into the induction chamber. Turn on the vaporizer at an induction level of 4% for isoflurane and 4 L/min for oxygen. After the animal is fully anesthetized, maintain the anesthetic with the nose cone and the anesthetic delivery at a level of 1.5% for isoflurane and 0.4 L/min for oxygen during surgery. Monitor the animal for respiration.</li>
		<li>Apply chlortetracycline hydrochloride eye ointment to prevent corneal dryness during the surgery.</li>
		<li>Shave the surgical area on the dorsal surface from the lower thoracic region to the top of the sacral region using a small animal trimmer. Remove the shaved fur with tissue wipes.</li>
		<li>Apply depilatory cream onto the shaved area and leave it there no longer than 3 min. Remove the cream with gauze and flush with 2 mL of 0.9% sterile saline.</li>
		<li>Place a custom-made surgical cylindrical pad (Figure 2A) under the abdomen of the mouse to raise up the lumbar spine and facilitate the surgical operation.</li>
	</ol>
	</li>
</ol>
2. Exposure of the lumbar third to lumbar fifth (L3&#8211;L5) spinous processes
<ol>
	<li>Use the index finger to touch the subcutaneous spinous processes of the lumbar vertebrae, which are more outward, and compare with thoracic vertebrae and sacral vertebrae to identify the lumbar region.</li>
	<li>Rinse the skin using 75% alcohol. Perform a 3&#8211;4 cm midline skin incision over the lumber region from the mid-thoracic region to the hip using a scalpel blade to expose the fascia.</li>
	<li>Identify the lumbar spine by the morphology of the posterior fascia inserted onto the tips of the spinous processes. In detail, the third lumbar (L3) to the first sacral (S1) fasciae are distinct from other fasciae by their &#8220;V&#8221; shapes. The last &#8220;V&#8221; tip connects to the first sacral (S1) fascia and the first &#8220;V&#8221; tip corresponds to the L3 spinous process (Figure 2B).</li>
	<li>Make the posterior paraspinous muscle incisions along the spinous processes from L3 to L5 on both sides laterally with a scalpel blade (Figure 2C). Control the incision depth towards the facets to reduce hemorrhage.</li>
	<li>Separate the muscle layers using two ophthalmic forceps to expose L3 to L5 spinous processes and supraspinous ligaments.</li>
</ol>
3. Resection of L3&#8211;L5 spinous processes along with the ligaments
<ol>
	<li>Separate individual spinous processes by cutting off interspinous ligaments using Venus shears (Figure 2D).</li>
	<li>Resect the L3&#8211;L5 spinous processes along with the interspinous ligaments with Venus shears (Figure 2E).</li>
	<li>Suture the skin incision with sterile silk braided (suture size 5.0) without reattachment of the paravertebral muscles.</li>
	<li>Apply Chlortetracycline Hydrochloride Eye Ointment to the surgical site.</li>
	<li>Administer Buprenorphine-SR (25 uL per gram of mouse weight) immediately after LSI surgery for analgesia.</li>
	<li>Place the animals in a warm chamber and monitor during recovery from the anesthesia. Monitor food and water intake before returning the animals to the home cage.</li>
	<li>Monitor the animal once daily for the first 3 days after operation. The animal should be able to have a normal appetite and should heal with no sign of pus, hemorrhage, or swelling. They may have minor impairment in locomotion.</li>
	<li>Carry out sham operations only by the detachment of the posterior paravertebral muscles from the L3&#8211;L5 vertebrae.</li>
</ol>

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The authors have nothing to disclose.

We developed the lumbar spine instability mouse model based on the cervical spondylosis mouse model in which the posterior paravertebral muscles from the vertebrae were detached and the spinous processes along with the supraspinous and interspinous ligaments were resected25. We performed a similar operation onto the lumbar spine, which has more prominent spinous processes. The LSI mouse model developed similar IDD in the lumbar spine.
The advantages of the LSI model include strong operability, no requirement of a special equipment, reproducibility, and a relatively short period of IDD development.
Some key points are presented here to help improve the success rate during the operation. These are also the critical steps. First, remove the hair on the lower back portion, as clear as possible, because any shaved hair left in the wound may cause anaphylactic reaction. Second, a cylindrical pad or any other pad is recommended to raise up the lumbar vertebrae. Third, use micro scissors to control the incision depth and hemorrhage. When hematocoelia is noticed during the operation, stop the operation and sacrifice the mouse since the mouse will not survive during or post-surgery. Fourth, reattachment of paraspinous muscle is not recommended because the reattachment may make up the instability. Fifth, a full resection of the whole L3&#8211;L5 spinous processes reduces variability in individual model. Sixth, avoid injuring surrounding nerves and blood vessels, otherwise, the mouse may develop non-canonic pathological changes. If the models do not exhibit the typical phenotype as shown in the results, check the above six points.
The success of this LSI model can be assessed by two golden standards including decreased IVD volume measured by small animal MRI or by &#956;CT, and an IVD score based on the histological observation. The LSI model develops IDD as early as 2 weeks post LSI surgery, but develops porosity in endplate as early as 1 week, as observed. It is suitable for study on nucleus pulpous shrinkage, endplate sclerosis, IDD related to osteoclast-induced cytokines, IDD-induced osteoporosis (16 weeks post LSI) etc.
There are some limitations in the LSI model. LSI operation is a relatively great trauma for the mouse. The inflammation is inevitable and usually seen within 7 days post operation. Thus, this model is not suitable for observing early pathological changes of IDD, especially within 7 days caused by mechanical loading changes.
The model can be modified by targeting on different lumbar vertebrae such as L5 only or from L1 to L5. Healthy control is also recommended in addition to sham groups.
In summary, we developed a surgical-induced lumbar IDD mouse model and have the procedure visualized to help others reproduce the animal model and apply it in IDD studies.

Intervertebral disc degeneration (IDD) is commonly seen in aging and even young people caused by many factors1. Surgery for patients who suffer from IDD, causing low back pain and impaired movement, is usually performed at a later stage or in severe cases and has potential risks such as nonunion or infection2. Ideal non-operative treatment requires comprehensive understanding of the IDD mechanism. The IDD animal model serves as a crucial tool for studies of IDD mechanism and evaluation of IDD treatment.
Larger animals have been chosen for IDD models such as primates, sheep, goats, dogs, and rabbits due to their similarity with human anatomical structure to a great extent and the strong operability in terms of size of intervertebral discs (IVDs)3,4,5,6,7,8. However, these animal models are time-consuming and cost-intensive9. Mouse IVD is a poor representation of the human IVD based on geometrical measurements of the aspect ratio, nucleus pulposus to disc area ratio, and normalized height10. Despite the difference in size, mouse lumbar IVD segment exhibits mechanical properties similar to human IVD such as compression and torsion stiffness11. In addition, mouse IDD model has the advantage of low cost, relatively short IDD development, and more options for genetically modified animals and antibodies utilized in further mechanistic studies12,13,14,15.
Experimental-induced IDD models vary from the inducers and applications. For example, collagenase-induced extracellular matrix (ECM) degeneration is appropriate for ECM regeneration research16. Genetically modified phenotype are suitable for studying the gene function in the IDD process and in genetic therapies17. Annulus fibrosus incision and smoke models mimic trauma and non-inflammation induced IDD12,18.
Spinal instability (SI) leads to an unstable spine that is not in an optimal state of equilibrium. It can be caused by abnormal movement of a lumbar motion segment due to the weakness of the surrounding supportive tissue such as ligaments and muscles. It is also commonly seen post spinal fusion operation19. SI is considered as the main cause of IDD. Therefore, we aim to develop a SI mice model (focused on lumbar spine) that mimics the human IDD process20,21.
In the protocol, we introduced the procedure of establishing lumbar spinal instability (LSI) mouse model by the resection of lumbar third (L3) to lumbar fifth (L5) spinous processes along with the supraspinous and interspinous ligaments (Figure 1A,B). The animal model develops IDD as early as 1-week post-surgery as shown by hypertrophy and porosity in endplates (EPs). IVD volume starts to decrease 2 weeks post-surgery through 16 weeks along with increased IVD score, which indicates the degree of IDD. We believe the detailed and visualized procedure is useful for researchers to establish the LSI mouse model in their laboratory and apply to IDD research as needed.

<table><tbody><tr><td>Chlortetracycline Hydrochloride Eye Ointment</td><td>Shanghai General Pharmaceutical Co., Ltd.</td><td>H31021931</td><td>Prevent eye dry, Prevent wound infection</td></tr><tr><td>C57BL/6J male mice</td><td>Tian-jiang Pharmaceuticals Company (Jiangsu, CN)</td><td>SCXK2018-0004</td><td>Animal model</td></tr><tr><td>Disposable medical towel</td><td>Henan Huayu Medical Devices Co., Ltd.</td><td>20160090</td><td>Platform for surgical operation</td></tr><tr><td>Inhalant anesthesia equipment</td><td>MIDMARK</td><td>Matrx 3000</td><td>Anesthesia</td></tr><tr><td>Isoflurane</td><td>Shenzhen RWD Life Technology Co., Ltd.</td><td>1903715</td><td>Anesthesia</td></tr><tr><td>Lidocaine hydrochloride</td><td>Shandong Hualu Pharmaceutical Co., Ltd.</td><td>H37022839</td><td>Pain relief</td></tr><tr><td>Medical suture needle</td><td>Shanghai Pudong Jinhuan Medical Products Co., Ltd.</td><td>20S0401J</td><td>Suture skin</td></tr><tr><td>Ophthalmic forceps</td><td>Shanghai Medical Devices (Group) Co., Ltd. Surgical Instruments Factory</td><td>JD1050</td><td>Clip the skin</td></tr><tr><td>Ophthalmic scissors(10cm)</td><td>Shanghai Medical Devices (Group) Co., Ltd. Surgical Instruments Factory</td><td>Y00030</td><td>Skin incision</td></tr><tr><td>silk braided</td><td>Shanghai Pudong Jinhuan Medical Products Co., Ltd.</td><td>11V0820</td><td>Suture skin</td></tr><tr><td>Small animal trimmer</td><td>Shanghai Feike Electric Co., Ltd.</td><td>FC5910</td><td>Hair removal</td></tr><tr><td>Sterile surgical blades(12#)</td><td>Shanghai Pudong Jinhuan Medical Products Co., Ltd.</td><td>35T0707</td><td>Muscle incision</td></tr><tr><td>Veet hair removal cream</td><td>RECKITT BENCKISER (India) Ltd</td><td>NA</td><td>Hair removal</td></tr><tr><td>Venus shears</td><td>Mingren medical equipment</td><td>Length:12.5cm</td><td>Clip the muscle and spinous process</td></tr></tbody></table>

a mouse model of lumbar spine instability

Intervertebral disc degeneration (IDD) is a common pathological change leading to low back pain. Appropriate animal models are desired for understanding the pathological processes and evaluating new drugs. Here, we introduced a surgically induced lumbar spine instability (LSI) mouse model that develops IDD starting from 1 week post operation. In detail, the mouse under anesthesia was operated by low back skin incision, L3&#8211;L5 spinous processes exposure, detachment of paraspinous muscles, resection of processes and ligaments, and skin closure. L4&#8211;L5 IVDs were chosen for the observation. The LSI model develops lumbar IDD by porosity and hypertrophy in endplates at an early stage, decrease in intervertebral disc volume, shrinkage in nucleus pulposus at an intermediate stage, and bone loss in lumbar vertebrae (L5) at a later stage. The LSI mouse model has the advantages of strong operability, no requirement of special equipment, reproducibility, inexpensive, and relatively short period of IDD development. However, LSI operation is still a trauma that causes inflammation within the first week post operation. Thus, this animal model is suitable for study of lumbar IDD.

The LSI mouse model is applied in the studies of IDD mechanism, IDD treatment, endplate (EP) degeneration such as sclerosis, and sensory innervation in EP20,21,22,23. The LSI mouse develops IDD and EP degenerative changes, as identified, by decreased IVD volume and height, increased EP volume, and increased IVD and EP scores.
The dissected and fixed lower thoracic and lumbar spine were examined by high-resolution Micro Computed Tomography (μCT) as previously described20,21. The lower thoracic lumbar with ribs were included for the identification of L3–L5 vertebrae (Figure 3A). X-rays of the L3–L5 spine on a lateral view indicate existence and inexistence of spinous processes in Sham and LSI groups (Figure 3B). The results are clearer by 3D-reconstruction of L3–L5 spine on a left anterior oblique view (Figure 3C) and by transverse image of a L3–L5 vertebrae (Figure 3D).
Coronal images of the L4–L5 IVD were used to perform 3D histomorphometric analyses of IVD20 (Figure 4A). IVD volume is defined as the region of interest (ROI) covering the whole invisible space between L4 and L5 vertebrae. Parameter: TV (total tissue volume) was used for 3D structural analysis (Figure 4B). IVD volume significantly increased 1 week post-surgery and started to decrease from 2 weeks to 16 weeks post operation as observed in Figure 4C.
The height of IVD space varied from the anterior to the posterior (Figure 4E,G). LSI had a significant effect on the rear site. Thus, the posterior one-third coronal plane of IVD space was chosen for IVD height measurement (Figure 4D,E). IVD height decreased from 2 weeks to 16 weeks post-surgery (Figure 4F), which was consistent with the findings in IVD volume (Figure 4C).
Coronal images of the L4–L5 IVD space were applied to 3D histomorphometric analyses of both cranial and caudal endplates (Eps) (Figure 5A). Endplate (EP) volume is defined to cover visible bony plate close to the vertebrae (Figure 5A,B)21. The anterior one-fourth coronal plane of five consecutive images of cranial EP were used for 3D reconstruction (Figure 5C), which showed increased cavities within cranial EP in LSI mice (Figure 5D). The results were also indicated by an increased percentage of trabecular separation values which were greater or equal to 0.089 (Figure 5E). Meanwhile, EP volumes significantly increased post-surgery (Figure 5F). Caudal EPs exhibit similar phenotype by LSI (Figure 5G,H), indicating LSI leads to EP hypertrophy and an increase in cavities.
L5 vertebral bodies were reconstructed by drawing the outline of all transverse sections of each L5 vertebral body without accessories and converting all 2D images to a 3D model. The construction and analysis were done with commercial software (e.g., NRecon v1.6 and CTAn v1.9, respectively). The volumes of L5 vertebra slightly increase post-surgery but only have statistic difference between sham group and 16-week LSI group (Figure 6B). A significant decrease in BV/TV was also present 16 weeks post-surgery, indicating that LSI causes vertebral bone loss at a later stage (Figure 6A,C).
LSI induces IVD degeneration and EP degeneration as indicated by increased IVD and EP scores24 (Figure 7A,C). A reduction in intracellular vacuoles of nucleus pulposus cells was accelerated in LSI groups (Figure 7B). Cavities increase in LSI EPs (Figure 7D) accompanied with increased number of osteoclasts as indicated by Trap staining (Figure 7E,F).
The data was shown as mean ± s.d. Statistical significance was determined by a Student’s t-test. The level of significance was defined as p &lt; 0.05. All data analyses were performed using SPSS 15.0.
<img alt="Rodent spinal anatomy diagram showing vertebrae, intervertebral ligaments, and spinal processes." class="xfigimg" src="/files/ftp_upload/61722/61722fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Schematic of LSI mouse model. (A) Anatomy of L3–L5 vertebrae in the lower back area of the mouse. (B) Resection of spinous processes along with interspinous ligaments and supraspinous ligaments (marked pale). A red dotted line indicates a section plane. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Mouse spinal cord exposure; L3-L6 identification; procedural sequence; anatomical study." class="xfigimg" src="/files/ftp_upload/61722/61722fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Exposure of L3–L5 spinous processes and LSI operation. (A) A custom-made cylindrical pad is placed under the mouse’s abdomen. (B) Exposure of the lumbar fasciae and identification of L3 to S1 spinous processes by “V” shapes. (C) Lateral paraspinous muscle incisions on both sides of L3 to L5 spinous processes. (D) Exposure of individual spinous processes by cutting off interspinous ligaments. (E) Resection of L3–L5 spinous processes with inter- and supra-spinous ligaments. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Vertebral imaging comparison; X-ray and 3D reconstructions of lumbar spine, Sham vs LSI method." class="xfigimg" src="/files/ftp_upload/61722/61722fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: LSI identification by μCT. (A) Localization of L3–L5 vertebrae by ribs with thoracic vertebrae in X-rays. (B) X-rays on a lateral view and (C) 3D reconstruction on a left anterior oblique view of L3–L5 vertebrae in Sham and LSI groups. (D) Transverse plane of lumbar vertebra with the resection of the spinous process. (D) has been modified from Bian et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="L4-5 IVD volume analysis; A: MRI diagram, B: transverse view, C: graph, D-E: sectional view, F-G: progression comparison." class="xfigimg" src="/files/ftp_upload/61722/61722fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: LSI reduces IVD volume. (A) Consecutive images of the invisible space (Red) between L4 and L5 EPs are used for 3D reconstruction. (B) IVD volume is defined by the TV of (A). (C) Quantification of L4–L5 IVD volume at five timepoints post-operation. N = 8 per group. Data is shown as mean ± s.d. *p &lt; 0.05, **p &lt; 0.01 versus Sham. (D) Transverse plane and (E) mid-sagittal plane of lumbar vertebral bodies. Blue double-arrows indicate anteroposterior diameter. Yellow line indicates posterior 1/3 plane. (F) Reconstruction of cranial and caudal EPs using five consecutive images of posterior 1/3 coronal plane of L4–L5. Red indicates IVD space. (G) Mid-sagittal plane of L4–L5. (C) has been modified from Bian et al.20. (F,G) have been modified from Bian et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Bone analysis: MRI images, trabecular separation chart, and bone volume graph in lumbar study." class="xfigimg" src="/files/ftp_upload/61722/61722fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: LSI induces EP hypertrophy and porosity. (A) Coronal plane of L4–L5. Red dotted line indicates image of caudal EP used for 3D construction. (B) Reconstruction of caudal L4 and cranial L5. Blue cartoon indicates caudal EP of L4–L5. (C) Mid-sagittal plane of L4–L5. Blue double-arrows indicate anteroposterior diameter. Yellow line indicates anterior 1/4 plane. (D) Reconstruction of cranial EPs by five consecutive images of anterior 1/4 plane of L4–L5. (E,G) Percentage of trabecular separation distribution of cranial (E) and caudal (G) EPs obtained from μCT analyses. (F,H) Quantification of cranial (F) and caudal (H). L4–L5 EP volume in indicated timepoints. N = 8 per group. Data is shown as mean ± s.d.* p &lt; 0.05 versus. Sham. (D–H) have been modified from Bian et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="L5 vertebra bone analysis; micro-CT images and statistical graphs showing TV and BV/TV changes, Sham vs. LSI." class="xfigimg" src="/files/ftp_upload/61722/61722fig06.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6: LSI causes vertebral bone loss at late stage. (A) Reconstruction of L5 vertebral bodies in 16-week Sham and LSI groups. (B,C) Quantification of L5 vertebra TV (B) and BV/TV (C). N = 8 per group. Data is shown as mean ± s.d.* p &lt; 0.05, ** p &lt; 0.01 versus. Sham. (B) has been modified from Bian et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Bar charts and microscopy images showing tissue changes in sham vs. LSI groups over 8 weeks." class="xfigimg" src="/files/ftp_upload/61722/61722fig07.jpg" data-altupdatedat="1747911167000" data-alt="Figure 7"> 
Figure 7: LSI leads to IVD and EP degeneration. (A) IVD score in LSI or sham mice as an indication of IVD degeneration. (B) Representative images of Safranin O staining for NPs in L4–L5 IVD. White indicates vacuoles. Red indicates proteoglycan. (C) EP score in LSI or sham group as an indication of EP degeneration. (D) Representative images of Safranin O-Fast green staining for caudal L4–L5 EPs. Green/blue stains calcified cavities. (E) Representative images of trap staining for caudal L4–L5 EPs. Purple indicates Trap+. N = 6 per group. Data is shown as mean ± s.d.* p &lt; 0.05, ** p &lt; 0.01 versus. Sham. (F) Quantification of Trap+ osteoclasts in (E). (A,B) have been modified from Bian et al.20. (C–F) have been modified from Bian et al.21. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61722/61722fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Mouse Model of Lumbar Spine Instability at JoVE.com

A Mouse Model of Lumbar Spine Instability

We developed a lumbar intervertebral disc degeneration mouse model by resection of L3&#8211;L5 spinous processes along with supra- and inter-spinous ligaments and detachment of paraspinous muscles.

Intervertebral disc degeneration (IDD) is a common pathological change leading to low back pain. Appropriate animal models are desired for understanding ...

a-mouse-model-of-lumbar-spine-instability

Nanyang Technological University

Research

JoVE Journal

Biology

7.6K Views.  Shanghai University of Traditional Chinese Medicine. Dit is een operatie om het muismodel te induceren biedt een alternatief voor lumbale tussenwervelschijfdegeneratie gerelateerde studies. Deze robuuste LSI-modus vereist geen speciale apparatuur. Het is reproduceerbaar en het kan worden gebruikt om de tussenwervelschijfdegeneratie in een relatief korte tijd te induceren.Het beheersen van de incisiediepten en de bloeding en het uitvoeren voor resectie van elk interspinous proces zijn twee belangrijke kosten voor implantatie van ons succe...

Video: A Mouse Model of Lumbar Spine Instability

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.

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.

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

Medicine

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 Neonatal Mouse Spinal Cord Compression Injury Model

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal cord. A major current area of research is focused on the mechanisms of adaptive plasticity that underlie spontaneous or induced functional recovery following SCI. Spontaneous functional recovery is reported to be greater early in life, raising interesting questions about how adaptive plasticity changes as the spinal cord develops. To facilitate investigation of this dynamic, we have developed a SCI model in the neonatal mouse. The model has relevance for pediatric SCI, which is too little studied. Because neural plasticity in the adult involves some of the same mechanisms as neural plasticity in early life1, this model may potentially have some relevance also for adult SCI. Here we describe the entire procedure for generating a reproducible spinal cord compression (SCC) injury in the neonatal mouse as early as postnatal (P) day 1. SCC is achieved by performing a laminectomy at a given spinal level (here described at thoracic levels 9-11) and then using a modified Yasargil aneurysm mini-clip to rapidly compress and decompress the spinal cord. As previously described, the injured neonatal mice can be tested for behavioral deficits or sacrificed for ex vivo physiological analysis of synaptic connectivity using electrophysiological and high-throughput optical recording techniques1. Earlier and ongoing studies using behavioral and physiological assessment have demonstrated a dramatic, acute impairment of hindlimb motility followed by a complete functional recovery within 2 weeks, and the first evidence of changes in functional circuitry at the level of identified descending synaptic connections1.

This article describes a method for generating a reproducible spinal cord compression injury (SCI) in the neonatal mouse. The model provides an advantageous platform for studying mechanisms of adaptive plasticity that underlie spontaneous functional recovery.

Spinal cord injury (SCI) typically causes devastating neurological deficits, particularly through damage to fibers descending from the brain to the spinal ...

A Tissue Displacement-based Contusive Spinal Cord Injury Model in Mice

Producing a consistent and reproducible contusive spinal cord injury (SCI) is critical to minimizing behavioral and histological variabilities between experimental animals. Several contusive SCI models have been developed to produce injuries using different mechanisms. The severity of the SCI is based on the height that a given weight is dropped, the injury force, or the spinal cord displacement. In the current study, we introduce a novel mouse contusive SCI device, the Louisville Injury System Apparatus (LISA) impactor, which can create a displacement-based SCI with high injury velocity and accuracy. This system utilizes laser distance sensors combined with advanced software to produce graded and highly-reproducible injuries. We performed a contusive SCI at the 10th thoracic vertebral (T10) level in mice to demonstrate the step-by-step procedure. The model can also be applied to the cervical and lumbar spinal levels.

We introduce a tissue displacement-based contusive spinal cord injury model that can produce a consistent contusive spinal cord injury in adult mice.

Producing a consistent and reproducible contusive spinal cord injury (SCI) is critical to minimizing behavioral and histological variabilities between ...

In Vivo Mouse Model of Spinal Implant Infection

Spine implant infections portend poor outcomes as diagnosis is challenging and surgical eradication is at odds with mechanical spinal stability. The purpose of this method is to describe a novel mouse model of spinal implant infection (SII) that was created to provide an inexpensive, rapid, and accurate in vivo tool to test potential therapeutics and treatment strategies for spinal implant infections.
In this method, we present a model of posterior-approach spinal surgery in which a stainless-steel k-wire is transfixed into the L4 spinous process of 12-week old C57BL/6J wild-type mice and inoculated with 1 x 103 CFU of a bioluminescent strain of Staphylococcus aureus Xen36 bacteria. Mice are then longitudinally imaged for bioluminescence in vivo on post-operative days 0, 1, 3, 5, 7, 10, 14, 18, 21, 25, 28, and 35. Bioluminescence imaging (BLI) signals from a standardized field of view are quantified to measure in vivo bacterial burden.
To quantify bacteria adhering to implants and peri-implant tissue, mice are euthanized and the implant and surrounding soft tissue are harvested. Bacteria are detached from the implant by sonication, cultured overnight and then colony forming units (CFUs) are counted. The results acquired from this method include longitudinal bacterial counts as measured by in vivo S. aureus bioluminescence (mean maximum flux) and CFU counts following euthanasia.
While prior animal models of instrumented spine infection have involved invasive, ex vivo tissue analysis, the mouse model of SII presented in this paper leverages noninvasive, real time in vivo optical imaging of bioluminescent bacteria to replace static tissue study. Applications of the model are broad and may include utilizing alternative bioluminescent bacterial strains, incorporating other types of genetically engineered mice to contemporaneously study host immune response, and evaluating current or investigating new diagnostic and therapeutic modalities such as antibiotics or implant coatings.

The protocol describes a novel in vivo mouse model of spinal implant infection where a stainless-steel k-wire implant is infected with bioluminescent Staphylococcus aureus Xen36. Bacterial burden is monitored longitudinally with bioluminescent imaging and confirmed with colony forming unit counts after euthanasia.

Spine implant infections portend poor outcomes as diagnosis is challenging and surgical eradication is at odds with mechanical spinal stability. The ...

Establishing a Mouse Contusion Spinal Cord Injury Model Based on a Minimally Invasive Technique

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, thereby improving the reproducibility of the experiments.
These methods need two requirements to be fulfilled: clarity of the surgical anatomical pathway and simplicity and convenience of the laboratory device. Crucially for the operator, a clear anatomical pathway provides minimally invasive exposure, which avoids additional damage to the experimental animal during the surgical procedures and allows the animal to maintain a consistent and stable anatomical morphology during the experiment.
In this study, the use of a novel integrated platform called the SCI coaxial platform for spinal cord injury in small animals to expose the T9 level spinal cord in a minimally invasive way and stabilize and immobilize the vertebra of mice using a vertebral stabilizer is researched, and, finally, a coaxial gravity impactor is used to contuse the spinal cord of mice to approach different degrees of T9 spinal cord injury. Finally, histological results are provided as a reference for the readers.

Minimally invasive techniques and a simple laboratory device improve the reproducibility of the spinal cord injury model by reducing operative damage to the experimental animals and allowing anatomical morphology maintenance. The method is worthwhile because the reliable results and reproducible procedure facilitate investigations of the mechanisms of disease reparation.

Using minimally invasive methods to model spinal cord injury (SCI) can minimize behavioral and histological differences between experimental animals, ...

Neuroscience

A Spinal Cord Injury Contusion Device for Accurate Mouse Models

Automated Impactor for Contusive Spinal Cord Injury Model in Mice

Spinal cord injury (SCI) due to traumatic injuries such as car accidents and falls is associated with permanent spinal cord dysfunction. Creation of contusion models of spinal cord injury by impacting the spinal cord results in similar pathologies to most spinal cord injuries in clinical practice. Accurate, reproducible, and convenient animal models of spinal cord injury are essential for studying spinal cord injury. We present a novel automated spinal cord injury contusion device for mice, the Guangzhou Jinan University smart spinal cord injury system, that can produce spinal cord injury contusion models with accuracy, reproducibility, and convenience. The system accurately produces models of varying degrees of spinal cord injury via laser distance sensors combined with an automated mobile platform and advanced software. We used this system to create three levels of spinal cord injury mice models, determined their Basso mouse scale (BMS) scores, and performed behavioral as well as staining assays to demonstrate its accuracy and reproducibility. We show each step of the development of the injury models using this device, forming a standardized procedure. This method produces reproducible spinal cord injury contusion mice models and reduces human manipulation factors via convenient handling procedures. The developed animal model is reliable for studying spinal cord injury mechanisms and associated treatment approaches.

Author Spotlight: Insight Into Innovations in Spinal Cord Injury Research

Presented here is a novel automated spinal cord injury contusion device for mice, which can accurately produce spinal cord injury contusion models with varying degrees.

Spinal cord injury (SCI) due to traumatic injuries such as car accidents and falls is associated with permanent spinal cord dysfunction. Creation of ...

A Mouse Model of Ankle-Subtalar Complex Joint Instability

Ankle sprains are perhaps the most common sports injuries in daily life, often resulting in instability of the ankle-subtalar complex joint (ASCJ), and can eventually lead to post-traumatic osteoarthritis (PTOA) in the long term. However, due to the complexity of the injury mechanism and the clinical manifestations, such as ecchymosis, hematoma, or tenderness in the lateral foot, there is no clinical consensus on diagnosing and treating ASCJ instability. Since the musculoskeletal structure of the bones and ligaments of the mouse hindfoot is comparable to that of humans, an animal model of ASCJ instability in mice was established by the transection of ligaments around the ASCJ. The model was well-validated through a series of behavioral tests and histological analyses, including a balance beam test, a footprint analysis (an assessment of exercise level and balance ability in mice), a thermal nociception assessment (an assessment of foot sensory function in mice), micro-computed tomography (CT) scanning, and section staining of the articular cartilage (an assessment of articular cartilage damage and degeneration in mice). The successful establishment of a mouse model of ASCJ instability will provide a valuable reference for clinical research on the injury mechanism and result in better treatment options for ankle sprain.

The ankle-subtalar complex joint (ASCJ) is the core of the foot and plays a key role in balance control in daily activities. Sports injuries often lead to instability in this joint. Here, we describe a mouse model of ligament transection-induced instability of the ASCJ.

Ankle sprains are perhaps the most common sports injuries in daily life, often resulting in instability of the ankle-subtalar complex joint (ASCJ), and ...

Compression Testing of Mouse Vertebra for Determination of Mechanical Strength

Mouse Lumbar Vertebra Uniaxial Compression Testing with Embedding of the Loading Surface

There is increasing awareness that cortical and cancellous bone differ in regulating and responding to pharmaceutical therapies, hormone therapies, and other treatments for age-related bone loss. Three-point bending is a common method used to assess the influence of a treatment on the mid-diaphysis region of long bones, which is rich in cortical bone. Uniaxial compression testing of mouse vertebrae, though capable of assessing bones rich in cancellous bone, is less commonly performed due to technical challenges. Even less commonly performed is the pairing of three-point bending and compression testing to determine how a treatment may influence a long bone's mid-diaphysis region and a vertebral centrum similarly or differently. Here, we describe two procedures to make compression testing of mouse lumbar vertebrae a less challenging method to perform in parallel with three-point bending: first, a procedure to convert a three-point bending machine into a compression testing machine, and second, an embedding method for preparing a mouse lumbar vertebra loading surface.

Author Spotlight: Enhancing Small Animal Bone Compression Testing for Research

In this protocol, two approaches are described to make uniaxial compression testing of mouse lumbar vertebrae more attainable. First, the conversion of a three-point bending machine to a compression testing machine is described. Second, an embedding method for preparing the loading surface that uses bone cement is adapted for mouse lumbar vertebrae.

There is increasing awareness that cortical and cancellous bone differ in regulating and responding to pharmaceutical therapies, hormone therapies, and ...

Surgical Induction of Lumbar Spine Instability in Mouse Model: A Surgical Procedure for the Resection of the Lumbar Spinous Processes and Connected Ligaments in Murine Model

Source: Liu, S. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/61722">A Mouse Model of Lumbar Spine Instability.</a> J. Vis. Exp. (2021)
In this video, we describe the surgical procedure to induce lumbar spine instability in a mouse model by resecting the Lumbar L3 to L5 spinous process along with supraspinous and interspinous ligaments.

Source: Liu, S. et al. A Mouse Model of Lumbar Spine Instability. J. Vis. Exp. (2021)
In this video, we describe the surgical procedure to induce lumbar ...

A Mouse Model of Lumbar Spine Instability

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