In Vivo Telemetry to Record Long-Term Cardiovascular Parameters, Temperature, and Activity in Spinal Cord Injury Rat Models

Traumatic Spinal cord injury (SCI) impairs sensory and motor functions and disrupts cardiovascular and thermal regulation^1,2,3,4. Injuries at or above the T6 level disrupt supraspinal control of autonomic function, leading to severe alterations in various cardiovascular responses, including impaired baroreceptor reflexes, blood pressure dysregulation, heart rate variability (bradycardia or tachycardia), thermodysregulation, and decreased motor activity^{3,5,6,7,8,9,10}. These abnormalities are observed during both the acute and chronic phases of injury^2,3. Due to age-related changes in demographics over the years, cardiovascular dysfunction has become one of the leading causes of mortality and long-term morbidity in individuals with SCI^9,11. The acute phase of injury causes neurogenic shock, which is often characterized by hypotension and bradycardia due to sympathetic disruption⁹ but can also lead to hypertension in individuals with injury above T6 level^12,13. During the chronic phase, these individuals often experience autonomic dysreflexia (AD) and orthostatic hypotension(OH)⁵. AD is a life-threatening condition defined as a sudden episodic increase in systolic blood pressure (SBP) of 20-40 mmHg above baseline^14,15. Individuals may experience a range of symptoms, including a pounding headache, skin flushing, sweating, nasal congestion, piloerection, anxiety, and blurred vision^13,14,16. If untreated, AD can elevate SBP up to 300 mm Hg, potentially leading to seizures, stroke, myocardial infarction, or sudden death^17,18. OH is characterized by sudden reduction in blood pressure (BP) upon transitioning from a supine or seated position to standing, potentially resulting in dizziness, lightheadedness, or fainting. It is clinically defined as a sustained decrease in systolic blood pressure of at least 20 mmHg or diastolic blood pressure (DBP) of at least 10 mmHg within 3 minutes of standing^19,20. Thus, the chronic phase is characterized by both hypertension and hypotension episodes in individuals with severe-high thoracic SCI. A detailed understanding of cardiovascular physiology, core body temperature fluctuations, and activity in animal models is essential for designing targeted interventions to address autonomic dysfunction during both the acute and chronic phases following SCI.

The placement of a telemetric implant in the descending aorta enables efficient recording of the impact of SCI on systolic and diastolic blood pressure, mean arterial pressure (MAP), heart rate, core body temperature, and activity in rodent models of experimental SCI. Understanding the robust and sensitive methodology for telemetric implant placement is crucial for acquiring reliable physiological data following SCI. Telemetric techniques for recording blood pressure offer several advantages over non-telemetric techniques (e.g., the tail cuff technique). The advantages include continuous and real-time monitoring, reduced stress-induced artifacts, enhanced data accuracy and reproducibility, improved animal welfare, and suitability for small or sensitive models, such as disease models, pregnant animals, and neonates^21,22,23. Owing to these benefits, telemetric techniques facilitate experimental reproducibility and enhance translational relevance in SCI research by enabling the modeling of human-like autonomic and neural responses in unrestrained, freely moving animals.

The present manuscript provides a detailed, step-by-step methodology for telemetric implant placement in the descending aorta²⁴, followed by a severe high-thoracic contusion SCI procedure^25,26. It demonstrates how various cardiovascular, core body temperature, and behavioral parameters can be recorded using Ponemah software. The manuscript also outlines the challenges associated with the procedure and offers potential solutions. Additionally, it highlights the various applications of in vivo telemetry in SCI research.

Ethical approval was obtained from the University of Kentucky Institutional Animal Care and Use Committee, and the procedures were followed strictly, adhering to the protocol guidelines. Adult female Wistar rats (11-13 weeks, 250-300 g, n = 3 ) were housed at the Division of Lab Animal Research animal facility (University of Kentucky, USA) under standard housing conditions (12/12 light: dark cycle, humidity 30%-70%, temperature 22 ± 2 °C) with food and water ad libitum. The reagents and the equipment used are listed in the Table of Materials.

1. Surgical Procedure for telemetric implant placement

NOTE: The placement of the telemetric implant was adapted from Rabchevsky et al. and further optimized to minimize complications and mortality²⁴. The duration of anesthesia and pre-surgical care is approximately 10 minutes. The telemetric implantation surgical procedure varies from 15-30 min. Abdominal and skin incision closure and post-surgical care take 15-20 min. The total duration of the telemetric implant placement surgical procedure is 40-60 min. The reagents and the equipment used for telemetric implantation surgery are listed in the Table of Materials.

1. Pre-surgery and surgical table preparations
   1. Autoclave surgical instruments, 16-ply gauze sponges, cotton-tip applicators, and occlusion threads. Fill the pressure catheter tip of the telemetric implant with gel using a syringe.
   2. Sterilize the implants by sequentially immersing them for 24 h each in detergent, cold sterilant, and 0.9% sodium chloride.
   3. Keep the implant submerged in 0.9% sodium chloride in a tube labeled with the implant number.
   4. Place a heating pad on the surgical table and set it to 37-38 °C. Cover the heating pad with surgical underpads, followed by sterile drape towels, to maintain a sterile field during the procedure.
2. Anesthesia and pre-surgical care
   1. Anesthetize the rat with isoflurane (3%-4%) and maintain anesthesia at 2% using a Low-Flow Electronic Vaporizer.
   2. Administer buprenorphine (0.02 mg/kg; analgesic), meloxicam (1 mg/kg; anti-inflammatory), and 5 mL of 0.9% normal saline subcutaneously prior to surgery. Apply lubricating gel to both eyes to prevent corneal dehydration.
3. Surgical site preparation, incision, and descending aorta exposure
   1. Shave the abdominal area from 1 cm above the urethral orifice to the mid-abdomen using an electric shaver. Clean the exposed skin three times with alcohol prep pads, followed by application of the antiseptic povidone-iodine. Place a press-and-seal barrier on the animal and cut a small opening to expose the surgical site.
   2. Make a 4.5-5 cm midline skin incision starting 1-1.5 cm above the urethral orifice, then perform an abdominal incision. Secure the visceral organs with sterilized, saline-soaked 16-ply gauze sponges. Use metal retractors to further secure the organs and to maintain visibility of the region to be operated on.
   3. Use two fine forceps (microdissection tweezers and mirror-finish forceps) to gently separate the fat bodies and expose the descending aorta and vena cava. Keep one forceps stable to hold the fat bodies, and use the other one to dissect.
4. Blunt dissection and occlusion
   1. Perform blunt dissection using the forceps mentioned above, approximately 0.5 cm below the renal artery (used as a landmark) to separate the descending aorta from the vena cava. In addition, separate the descending aorta from the underlying fat bodies to create space for angled forceps insertion.
   2. Insert angled forceps under the descending aorta through the dissected space between the aorta and the vena cava to ensure that the occlusion thread can pass through.
   3. Pass a sterile 4-0 silk thread underneath the descending aorta through the dissected space, and secure both ends of the thread using a hemostat.
   4. Have another experimenter hold the hemostat at approximately a 45-degree angle, make angular adjustments to minimize or eliminate blood backflow.
   5. Temporarily mono-occlude the descending aorta proximal to the catheter insertion point, at the point of separation of the descending aorta and vena cava
5. Insertion and securing of the implant
   1. Remove the implant from 0.9% sodium chloride, inspect the pressure catheter tip for air bubbles or gaps, and add gel if any of these are present before insertion.
   2. Pierce the upper wall of the descending aorta using a twisted 21-G needle, as demonstrated in the video, and insert the pressure catheter tip (diameter of tip 0.74 mm) of the telemetric implant. If the pressure catheter tip does not enter the aorta, guide it using a vein pick at the puncturing site.
   3. Insert the catheter tip 1.5-2 cm into the descending aorta. Clean any blood around the insertion site and apply a few drops of tissue adhesive to secure the implant and prevent bleeding. Check for any blood leakage before proceeding forward.
   4. Anchor the implant body to the abdominal wall subcutaneously, slightly away from the incision site, by suturing the suture rib with non-absorbable sutures to ensure stability throughout the experiment.
6. Abdominal wall and incision closure
   1. Close the abdominal wall using absorbable sutures in an interrupted pattern with square knots. Close the skin using an intradermal, continuous suturing technique.
   2. Clean the incision area with hydrogen peroxide to remove any blood residue. Follow with an application of povidone-iodine solution, then apply a triple antibiotic cream containing Polymyxin B Sulfate, Bacitracin Zinc, and Neomycin Sulfate to facilitate healing.
7. Post-surgical care
   1. Immediately after surgery, inject the antibiotic enrofloxacin (5 mg/kg) subcutaneously. Place the rat in a home cage with a heating pad underneath, positioned supine, until it recovers from anesthesia.
      NOTE: Most animals recover from anesthesia immediately, but some may take 5-10 min.
   2. Once the rat is awake, fit an e-collar for 3 days around its neck to prevent biting of the abdominal sutures.
   3. For thermoregulation, keep the animal on alpha pads in a recovery cage with a water heating pad at 37-38 °C underneath for at least 24-72 h. Position half of the cage on the heating pad, allowing animals to move away from the heat if needed.
   4. Administer post-operative medications subcutaneously: buprenorphine and meloxicam (twice daily for 3 days), and enrofloxacin (once daily for 5 days) to manage pain, inflammation, and infection. Provide food and water ad libitum.

2. Surgical procedure for thoracic (T3) contusion injury

NOTE: The T3 contusion procedure was adapted from Squair et al.²⁶. The total duration of the T3 contusion injury procedure is approximately 40-45 min. The materials required for the T3 contusion SCI are listed in the Table of Materials. Perform the T3 contusion at least two weeks after the telemetric implant placement. The 2-week window was optimized based on observations that most animals recover from surgical distress and return to baseline blood pressure within 7-10 days. However, some animals may take a little longer.

1. Pre-surgery preparations
   1. Anesthetize the rat with intraperitoneal ketamine (80 mg/kg) and xylazine (7 mg/kg). Administer buprenorphine (0.02 mg/kg), meloxicam (1 mg/kg), and 5mL of 0.9% normal saline subcutaneously prior to surgery.
   2. Shave the dorsal surface from the cervical to mid-thoracic region using an electric shaver. Clean the skin three times with alcohol prep pads, followed by application of 5% povidone-iodine solution. Use lubricating eye gel to prevent corneal dehydration.
   3. Place the rat on a heating pad with surgical underpads covered with sterile drape towels, as done during the telemetric surgical procedure.
2. Incision, blunt dissection, and laminectomy
   1. Identify the T2 spinous process by its prominent dorsal spine and mark it. Make a dorsal midline incision from C8 to T5 and secure the area with muscle retractors.
   2. Perform blunt dissection using fine, small scissors and a curette through the superficial and deep paraspinal musculature. Avoid venous corona mortis around T4/T5 during dissection, as damaging it can cause animal's immediate death.
   3. Perform a T3 laminectomy to expose the T3 spinal segment.
3. Spinal cord contusion
   1. Secure the animal at the T2 and T4 spinal processes using Allis forceps on the IH impactor stage.
   2. Align the impactor tip over the midline using the 3D coordinate system to ensure an accurate midline hit. Set the parameters as required (Force = 400 kdyn with 5 s dwell time).
4. Incision closure and post-surgical care
   1. Suture the paraspinal muscles with absorbable sutures and seal the incision site with staples. Remove any blood residues near the incision area using hydrogen peroxide.
   2. Proceed with an application of povidone-iodine solution, then smear a triple antibiotic cream containing Polymyxin B Sulfate, Bacitracin Zinc, and Neomycin Sulfate to facilitate healing. Administer enrofloxacin (5 mg/kg) subcutaneously, immediately after the injury. Place the rat on alpha pads in recovery cages with heating pads underneath.
   3. Provide post-operative care, including buprenorphine and meloxicam (twice daily for 3 days), and enrofloxacin (once daily for 5 days). Manually express the bladder (Crede maneuver). Provide food and water ad libitum.

3. Settings for recording and exporting data

NOTE: The settings for recording and exporting cardiovascular, core body temperature, animal activity parameters, pre- and post-SCI using Ponemah software (version 6.51) are mentioned.

1. Data recording
   1. For recording the various parameters, click on Create under the Experiment tab and label the experiment.
   2. Go to the Hardware tab, click on Edit APR Configuration, select the APR, and click on Add to move it to the available region. Then, click on Edit PhysioTel/HD MX2 Configuration in the hardware tab to select the MX-2, click on Add, and confirm that it appears in the left panel. Add implants assigned to different animals from the implant inventory on the right side to the middle and left panels. The number of implants that can be added depends on the number of receiver plates connected with the system. Click on each added implant (labeled with animal number) in the left panel and connect it to a different receiver plate. Save and exit once all connections are made.
   3. Go to the Set up tab and then choose Experiment Setup, click on Enable Page, then click on the black background. Choose the subject, add pressure in the label, change the unit to mmHg, set the low and high values, and choose a color for displaying blood pressure traces.
   4. Go to the Set up tab and then choose Animal Setup. Click on the animal number, select the gender and species, then click on Pressure to enable parameters. If heart rate needs to be displayed, click on Heart Rate. Apply channel settings to similar channels, then click OK. This opens a window with different implants arranged in vertical order.
   5. Click on All Continuous on the top to visualize the blood pressure recording traces on the screen.
2. Data exporting
   1. Begin by navigating to the Experiment tab, clicking on Open, and selecting the data from the intended folder.
   2. Go to the Action tab, click on Start Review, and select the subject number, desired signal types, and time range by checking the intended outcomes. Note that the software exports up to 3GB of data at a time. The data can be derived by setting the logging rate in the actions tab, which determines how the data is segmented (in seconds, minutes, or hours).
   3. Once selections are made, click on the Experiment tab, then choose Save Marked Sections followed by Save Derived Data.
   4. Finally, go to the Actions tab and click on Close Review Session to end the data export.

The placement of a telemetric implant in the descending aorta allows measurement of autonomic dysreflexia and various cardiovascular parameters, including systolic blood pressure (SBP), diastolic blood pressure (DBP), mean arterial pressure (MAP), and heart rate (HR), pre- and post-SCI. The implant also measures core body temperature and activity. Comparing pre-injury and post-injury data reveals how cardiovascular outcomes, temperature, and activity are affected after SCI. Figure 1 presents the experimental design (Figure 1A-C) and representative outcomes, including autonomic dysreflexia (Figure 1D), cardiovascular parameters, including systolic blood pressure (SBP; Figure 1E (i)), diastolic blood pressure (DBP; Figure 1E (ii)), mean arterial pressure (MAP; Figure 1E (iii)), and heart rate (HR; Figure 1E (iv)), core body temperature (Figure 1E (v)), and animal activity (Figure 1E (vi)), pre- and post-spinal cord injury (Figure 1E). To minimize animal use, the data in Figure 1 were taken from an ongoing study involving female rats; however, the same procedure can be applied to male rats.

The representative graph in Figure 1D shows systolic blood pressure 1 min before the stimulus, during the stimulus, and 1 min after stimulus application in rats with T3 contusion injury after 8 weeks post-SCI using colorectal distension (CRD). CRD utilizes insertion of a balloon catheter into the rat's anus to stimulate the rectum24. The balloon is inflated once the baseline stabilizes, and the stimulus is applied for 1 min. An increase of at least 20 mm Hg SBP during stimulation over baseline SBP (prior to the stimulation) shows development of AD in the chronic phase of injury (8 weeks).

The graphs in Figure 1E compare recordings taken 48 h before T3 contusion injury and during the first two days post-injury (48 h after injury). After SCI, graphs show elevated blood pressure indicated by increased SBP, DBP, and MAP (Figure 1E (i), (ii), (iii)). Post-SCI results showed increased heart rate fluctuations (Figure 1E (iv)), irregularities in core body temperature (Figure 1E (v)), and reduced activity (Figure 1E (vi)). Additionally, SCI disrupted day-night rhythms in SBP, DBP, MAP, HR, core body temperature, and activity.

Figure 1: Schematic of telemetry implant placement and applications following a severe-high thoracic (T3) contusion injury. Insertion of an in vivo telemetric implant into the descending aorta of a rat (A), followed by severe high thoracic (T3) contusion injury using an IH impactor (Force = 400 kdyn with 5-s dwell time) 2 weeks later (B). Representative setup of telemetric recording using a computerized monitoring system with Ponemah software (C). Representative trace (data acquired with a logging rate of 2 s) of female rats showing autonomic dysreflexia (AD) 8 weeks after spinal cord injury (SCI). The green dotted line indicates a 1 min colorectal distension (CRD) stimulus (D), and a representative hourly mean for 48 h. i. systolic blood pressure (SBP), ii. diastolic blood pressure (DBP), iii. mean arterial pressure (MAP), iv. heart rate (HR), v. core body temperature (core body temp), and vi. activity pre- and post-SCI (E). The unshaded and shaded regions represent 12-h light and 12-h dark cycles, respectively. "0" indicates lights on, and "12" indicates lights off. The error bars represent mean ± SEM (n = 3). Please click here to view a larger version of this figure.

Placing a telemetric implant in the descending aorta provides a reliable method for measuring cardiovascular parameters, core body temperature, and animal activity. Telemetry offers clear advantages over non-telemetric techniques. It allows data collection from awake, freely moving animals, eliminating the confounding effects of anesthesia, which can alter blood pressure and heart rate^21,27. Unlike non-telemetric methods, telemetry prevents thermal and animal restraining stress, ensuring accurate cardiovascular measurements^23,28. It enables continuous monitoring of blood pressure, heart rate, core body temperature, and activity for weeks or even months, whereas non-telemetric procedures record only intermittently^21,28,29. Overall, telemetric monitoring provides greater accuracy and precision than non-telemetric approaches²⁷.

Telemetric methods diminish physical and physiological stress, preserving natural behavior and ensuring compliance with animal welfare guidelines²⁷. Their high precision and lower variability compared with non-telemetric techniques decrease the number of animals required and increase statistical power^21,23. Telemetry also offers strong translational value in drug discovery and cardiovascular research, as the data closely resemble human physiological responses²⁸. Collectively, the telemetric approach is superior and ethically preferable to non-telemetric methods.

Telemetric implant placement in the descending aorta of rats offers several advantages for SCI research compared with placement in the carotid or femoral arteries, which serve as alternative implant sites in animal models. Being a large central vessel, the descending aorta provides stable and representative measurements of blood pressure and heart rate, critical indicators for assessing cardiovascular dysfunction after SCI^30,31. Implantation at this site minimizes interference with limb blood flow and reduces the risk of ischemia and other complications often associated with peripheral artery placement^30,31. It also allows continuous monitoring of basal cardiovascular parameters and autonomic dysreflexia, both crucial in SCI research²⁴. Implanting in the femoral artery may compromise blood supply to the ipsilateral hindlimb, potentially affecting experimental outcomes and animal welfare^32,33. Although technically simpler, carotid artery implantation carries risks such as stroke or neurological complications due to its proximity to the cerebral circulation, which is undesirable in studies focused on peripheral cardiovascular effects³³. Therefore, implantation in the descending aorta provides a dependable site, minimizes local complications, and enhances both the experimental rigor and translational value of data collected from conscious, freely moving animals. Several common challenges are encountered while placing telemetric implants in the descending aorta.

Through trial and error, the procedure was optimized to overcome the following obstacles: (1) Abdominal inflammation: Abdominal inflammation was observed in rats at both acute and chronic stages following telemetric implant surgery. To mitigate this, a post-operative regimen consisting of the analgesic buprenorphine and anti-inflammatory meloxicam was implemented, resulting in a reduced incidence of abdominal inflammation. Previously, in each cohort, a few animals developed bloated, firm abdomens and eventually died. During implant retrieval, the abdominal cavity often contained clear fluid or, in some cases, brownish fluid with a pungent odor. This regimen reduced the rate of inflammation by approximately 70%-80% compared to animals that underwent surgery without it; (2) Extensive bleeding during dissection of the descending aorta from the vena cava: Use fine forceps (microdissection tweezers and mirror finish forceps) to gently separate the descending aorta from the vena cava using blunt dissection (hold and tear). Avoid cutting between the two using scissors or a blade, as it can cause extensive bleeding due to rupture of the underlying vasculature; (3) Insertion of telemetric implant in superficial layer: During descending aortic piercing, ensure that the needle passes through all three layers of the upper wall and not just the superficial layer. Oozing of blood after piercing indicates that all layers of the upper wall are pierced. The telemetric implant will not record properly if the pressure catheter is inserted only in the superficial layer; (4) Insertion of incorrect catheter length: Use a microscope during and/or after implant insertion to ensure the pressure catheter crosses the occlusion thread. The implant will not function properly if the inserted catheter length is incorrect; (5) Blood leakage from descending aorta: Allow 30-45 s after implant insertion for the adhesive to dry. This ensures the piercing in the descending aorta is fully sealed and not bleeding. Reseal if minor bleeding occurs due to blood backflow. Avoid pulling the implant's pressure catheter tip while suturing the implant body to the side of the abdominal wall, as this can cause bleeding; (6) Rats removing abdominal sutures: Rats may bite and remove sutures, leading to skin and internal organ damage. To reduce this, use uninterrupted sutures with a square knot for closing the abdominal wall. For skin closure, apply intradermal continuous sutures to minimize the animal's access to the sutures and knots. Apply an e-collar for 3 days postoperatively to further restrict access. Ensure that the fit is snug but not restrictive by placing an index finger between the collar and the rat's neck; (7) Hindlimb ischemia: Previously, the aorta was occluded at two locations: one at the descending aorta 0.5 cm below the renal artery, and another above the iliac bifurcation. The bi-occlusion technique helped to keep the artery taut, facilitating smoother implant placement and limiting blood backflow. However, it led to hindlimb ischemia in some animals if catheter insertion is not performed rapidly. Presently, a mono-occlusion 0.5 cm below the renal artery is performed, which effectively eliminates hindlimb ischemia in rats. With bi-occlusion, approximately 20%-30% ischemia was observed, whereas mono-occlusion completely eliminated hindlimb ischemia.

Applications

Telemetric implants (HD-S10) are magnetically activated devices designed to record various physiological parameters in small animal models. The placement of these implants in the descending aorta is a precise and dependable method for continuous hemodynamic monitoring in unrestrained, unanesthetized, and freely moving animals. This methodology allows preclinical researchers to perform real-time assessments of various cardiovascular parameters, temperature, and animal activity in the context of disease modeling, such as SCI, drug efficacy evaluation, and diurnal changes in autonomic function parameters. Researchers can acquire comprehensive physiological data from a single telemetric implant in an animal, covering both acute and chronic timepoints, making it an efficient technique. Figure 1 highlights this telemetry application and others currently used in the laboratory.

Hemodynamic parameters, including SBP, DBP, and MAP, can be recorded over both acute and chronic periods. Monitoring the 24/7 rest-activity rhythm can serve as a reliable marker of the acute effects of SCI³⁴. Simultaneous heart rate monitoring enables the detection of heart rate variability, including tachycardia and bradycardia. Comparing blood pressure and heart rate fluctuations pre- and post-SCI allows researchers to understand how cardiovascular physiology changes following injury. A robust daily rhythm in cardiovascular parameters, core body temperature, and activity is important for maintaining physiological homeostasis and overall health. Changes in diurnal core body temperature are also indicative of impaired thermoregulation after SCI³⁴. It is well established that SCI is a whole-body syndrome. Therefore, incorporating telemetric-based outcomes allows researchers to collect comprehensive activity data post-SCI beyond locomotor recovery, which is typically evaluated in isolation.

However, there are certain limitations of this technique that need to be considered. The placement of a telemetric probe in the descending aorta is a complicated procedure and therefore requires microsurgical expertise. The implant can become dislodged later due to various reasons, including mishandling during bladder expression in SCI rats, infection, tissue irritation, or internal clot formation. The cardiovascular recording will be affected if any of these occur; the data should be excluded. In addition, if a rat does not recover properly, the baseline cardiovascular data may be compromised. The telemetric setup and implants are expensive and require specialized software and equipment for data acquisition. Severe SCI may possibly lead to increased mortality; therefore, careful planning of animal numbers is necessary to maintain the overall statistical power of the study.

Overall, assessing cardiovascular dysfunctions, core body temperature, and activity using telemetry after SCI, across acute and chronic periods, is critical for understanding the impact of injury on cardiovascular health, overall well-being, and for designing potential interventions.