Anesthetized mice exhibit non-physiological systemic blood pressure, which precludes meaningful assessment of autonomic tone given the intimate relationship between blood pressure and the autonomic nervous system. Thus, a novel method to simultaneously record renal sympathetic nerve activity and blood pressure with intravenous infusion in conscious mice is outlined.
Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity (RSNA) is of great interest in many areas of research such as chronic kidney disease, hypertension, heart failure, diabetes and obesity. Unequivocal assessment of the role of the sympathetic nervous system is thus imperative for proper interpretation of experimental results and understanding of disease processes. RSNA has been traditionally measured in anesthetized rodents, including mice. However, mice usually exhibit very low systemic blood pressure and hemodynamic instability for several hours during anesthesia and surgery. Meaningful interpretation of RSNA is confounded by this non-physiological state, given the intimate relationship between sympathetic nervous tone and cardiovascular status. To address this limitation of traditional approaches, we developed a new method for measuring RSNA in conscious, freely-moving mice. Mice were chronically instrumented with radio-telemeters for continuous monitoring of blood pressure as well as a jugular venous infusion catheter and custom-designed bipolar electrode for direct recording of RSNA. Following a 48-72 hour recovery period, survival rate was 100% and all mice behaved normally. At this time-point, RSNA was successfully recorded in 80% of mice, with viable signals acquired up to 4 and 5 days post-surgery in 70% and 50% of mice, respectively. Physiological blood pressures were recorded in all mice (116±2 mmHg; n=10). Recorded RSNA increased with eating and grooming, as well-established in the literature. Furthermore, RSNA was validated by ganglionic blockade and modulation of blood pressure with pharmacological agents. Herein, an effective and manageable method for clear recording of RSNA in conscious, freely-moving mice is described.
Interest in using mice in several areas of biomedical research continues to expand with the development of countless genetically engineered models. For the most part, technical advances have kept pace with the increased use of mice in physiology and there is now an impressive selection of miniaturized devices developed specifically for measuring important physiological parameters in mice. Although telemetric devices for direct measurement of autonomic nervous tone in the conscious rat have been available for over a decade, miniaturized devices for assessing nerve activity in conscious mice are currently not available. Investigators typically circumvent this limitation by evaluating the contribution of the autonomic nervous system with indirect methods (i.e. plasma or urine catecholamines, pharmacological autonomic blockade, spectral analysis of patterns of blood pressure/heart rate)1.
While these approaches provide valuable information, the result is a global picture of overall autonomic tone, rather than revealing the discrete contribution of isolated populations of nerves to the phenomenon under investigation. Alternatively, direct recording of activity from specific nerves has been executed in anesthetized mice, which poses a multitude of concerns. It is exceedingly difficult to maintain stable blood pressure within the physiological range in an anesthetized mouse for several hours following surgery. In fact, in these types of experiments, blood pressure is often unreported or presented at extremely low levels (i.e. 60-80 mmHg vs >100mmHg in a conscious mouse)2. The fragility of the cardiovascular system exhibited in an anesthetized mouse preparation often precludes meaningful assessment of autonomic nerve activity, given the codependent relationship between blood pressure and sympathetic tone3,4.
To address this limitation, a new method for direct recording of renal sympathetic nerve activity (RSNA) in conscious, unrestrained mice, undisturbed within their home cages was developed. Both the surgical and experimental approach for successful implementation of this technique is described in detail. This preparation enables the investigator to simultaneously record arterial pressure via radiotelemetry in addition to RSNA, with the added capability to intravenously infuse agents of interest without disturbing the mouse.
Twenty four hours post-surgery, mice behave normally and do not exhibit signs of pain or distress. Experimental recordings may then commence 48 to 72 hours post-surgery while the mouse rests comfortably in its home cage with unrestricted access to food, water and environmental enrichment. Clear RSNA traces are presented and the characteristic responses of this nerve population to normal physical movements of the animal (such as eating and grooming) are demonstrated in addition to pharmacological modulation of systemic blood pressure. The quality and specificity of the RSNA signal is further validated by ganglionic blockade. This manuscript includes the audiovisual complement to an initially published description of this technique5.
All of the experimental procedures are in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee of the University of Mississippi Medical Center.
1. Animals and Housing
2. Customized Fabrication of the Implantable RSNA Electrode
NOTE: Construct the implantable RSNA electrode at least a few days in advance of the scheduled surgical procedure to accommodate curing and sterilization time (described below).
3. Construction of the Electrode Tip
4. Fine Preparation of the Electrode Tip for Recording
5. Construction of the Anchoring Pedestal
6. Sterilization of the Completed Implantable Electrode
7. Anesthesia and Preparation for Surgery
8. Surgical Implantation of the RSNA Electrode
9. Implantation of Blood Pressure Radiotelemeter
10. Implantation and Exteriorization of the Jugular Venous Catheter
11. Securing Exteriorized Electrode Leads
12. Post-Surgical Recovery
13. Experimental Setup for Recording Blood Pressure and RSNA
14. Sample Experimental Protocol and Validation of RSNA Signal
15. Data Analysis
Following the described protocol, survival rate was 100% – all mice instrumented in this study survived and recovered well following the surgical procedure. Within 24 hours of surgical preparation, all mice behaved normally, exhibiting typical eating, grooming and exploratory behaviors. No animals showed any sign of pain or distress at this time. 48 hours following surgery, a verifiable and clear RSNA signal was recorded in 10 out of the 12 mice. This signal was maintained in these mice 72 hours post-surgery, however a true RSNA signal was recorded in 7 (70%) of mice by day 4 and in only 5 (50%) mice by day 5 post-surgery. Mice that did not exhibit a high-quality RSNA signal due to electrical noise or contamination by electrocardiogram signals were still in good health until the time of euthanization.
Mean arterial pressure in the conscious mice 48 hours post-surgery was 116±2 mmHg, with a corresponding mean heart rate of 596±22 bpm (n=10). Simultaneous recording of a representative sample of blood pressure and RSNA at this time demonstrated clearly visible and characteristically rhythmic bursts of RSNA (Figure 2). The typical increases of RSNA expected with normal activities such as eating and grooming, as directly observed and noted by personnel, were also present (Figure 3). High-quality RSNA was also recorded sequentially in 50% of the mice under investigation up to 5 days after surgical preparation (Figure 4). Blood pressure and heart rate remained stable for the 5 day investigation period and values were not different from those we have recorded following up to 10 days of post-surgical recovery (Table 1)8.
To validate the RSNA signal and verify that it is indeed entrained with the arterial baroreflex, blood pressure was pharmacologically manipulated with an intravenous injection of sodium nitroprusside and phenylephrine. RSNA characteristically increased in response to the sodium nitroprusside-induced reduction of arterial pressure; conversely, RSNA was virtually silenced following the phenylephrine-induced increase in arterial pressure (Figure 5). Quantitatively, sodium nitroprusside decreased blood pressure to 62±3 mmHg, which corresponded to an elevation of RSNA to 77±9% above baseline levels (n=5; P<0.05, Figure 6). Similarly, following phenylephrine administration, arterial pressure reached 137±6 mmHg, which reduced RSNA by 79±2% below baseline level (n=5; P<0.05, Figure 6). Furthermore, RSNA was completely eliminated following ganglionic blockade with hexamethonium (Figure 7), establishing the post-ganglionic nature of the RSNA signal.
Figure 1: Construction and placement of the implantable renal sympathetic nerve electrode. Schematic depiction of the design and recommended placement of the implantable renal sympathetic nerve electrode. (A) Bipolar leads fitted with pin connectors and a third ground wire. (B) Wires are threaded through polyethylene (PE) 90 tubing to protect the exteriorized leads. (C) Design of the electrode tip in order to separate the bipolar leads from the ground wire. (D) Electrode tip is bent at a 90° angle to facilitate optimal position; the renal nerve bundle is placed perpendicular to the bipolar leads and wax-based laboratory film insulates the leads from the ground wire that is in contact with the underlying tissue. Reproduced with permission5. Please click here to view a larger version of this figure.
Figure 2: Representative recording of arterial pressure and renal sympathetic nerve activity (RSNA). Sample trace demonstrating simultaneous recording of systemic arterial blood pressure, RSNA and integrated RSNA in a conscious, quietly resting mouse 48 hours following surgical preparation. Reproduced with permission5. Please click here to view a larger version of this figure.
Figure 3: Response of renal sympathetic nerve activity (RSNA) to normal physical activity. Representative trace featuring simultaneous recording of systemic arterial blood pressure, RSNA and integrated RSNA in two conscious mice 48 and 72 hours following surgery at baseline and (A) upon commencement of active grooming or (B) quiet eating. The large arrow denotes commencement of the physical activity from rest. Reproduced with permission5. Please click here to view a larger version of this figure.
Figure 4: Long-term renal sympathetic nerve activity (RSNA) signal viability. Sequential representative recordings of blood pressure and RSNA in one conscious, quietly resting mouse several days following surgical preparation. (A) 2 days, (B) 3 days, (C) 4 days and (D) 5 days post-surgery. Please click here to view a larger version of this figure.
Figure 5: Entrainment of the renal sympathetic nerve activity (RSNA) signal with the arterial baroreflex. Representative recording of arterial blood pressure and RSNA in a conscious mouse at rest during (A) baseline and after subsequent intravenous administration of (B) sodium nitroprusside followed by (C) phenylephrine. Reproduced with permission5. Please click here to view a larger version of this figure.
Figure 6: Quantitation of renal sympathetic responsivity to arterial blood pressure. Quantitative response of arterial blood pressure and renal sympathetic nerve activity (RSNA) to pharmacological manipulation with sodium nitroprusside and phenylephrine. (A) Mean arterial pressure at baseline (black bar; 116±2 mmHg) and following subsequent intravenous administration of sodium nitroprusside (grey bar; 62±3 mmHg) and phenylephrine (open bar; 137±6 mmHg). (B) Corresponding RSNA response during sodium nitroprusside (grey bar; 77±9%) or phenylephrine (open bar; -79±2%). RSNA is expressed a percent change from baseline, mean ± SEM. *Significant difference from baseline (p<0.05, n=5). Reproduced with permission5. Please click here to view a larger version of this figure.
Figure 7: Post-ganglionic nature of renal sympathetic nerve activity (RSNA). Representative trace of arterial blood pressure and RSNA at (A) baseline, (B) immediately following ganglionic blockade with hexamethonium and (C) post-mortem. Reproduced with permission5. Please click here to view a larger version of this figure.
Animal ID | 2d | 3d | 4d | 5d | |
A | mmHg | 112 | 110 | 108 | 109 |
bpm | 657 | 551 | 626 | 616 | |
B | mmHg | 115 | 107 | 111 | 110 |
bpm | 582 | 652 | 662 | 668 | |
C | mmHg | 115 | 118 | 113 | 111 |
bpm | 591 | 599 | 689 | 664 | |
D | mmHg | 114 | 115 | 116 | 110 |
bpm | 457 | 513 | 599 | 531 | |
E | mmHg | 109 | 109 | 103 | 105 |
bpm | 632 | 687 | 699 | 689 |
Table 1: Baseline mean arterial pressure and heart rate values in instrumented mice over 5 consecutive days following surgery. Reproduced with permission5.
Herein we have outlined, demonstrated and validated a novel method for targeted evaluation of RSNA in conscious mice, free to move and rest comfortably in their home cages. Following surgical implantation of an arterial pressure radiotelemeter, an indwelling intravenous infusion catheter and a custom-designed bipolar RSNA electrode, mice recovered from surgery and were left undisturbed for 48 to 72 hours. Mice remained comfortably settled in their home cage at all times (including experimental periods) with unrestricted access to food, water and environmental enrichment. All ensuing experimental manipulation by the investigator was remote and did not disquiet the animals. Regarding the quality and interpretation of the RSNA signal, this approach completely removed the undesirable and unavoidable physiological complications of anesthesia and surgical trauma as well as restraint and other sources of physical and mental stress to the animal. Thus, these serious confounding factors which invariably impact the interpretation of sympathetic nerve activity measurements were effectively eliminated.
All mice were in good health and as early as 24 hours post-surgery, displayed typical behaviors such as brightness, activity, responsiveness, eating, drinking, grooming as well as playful and exploratory behavior. All animals exhibited these characteristics and actively engaged with the provided environmental enrichment regardless of whether or not a viable RSNA signal was able to be recorded. Although the recovery time required to completely restore normal blood pressure following implantation of the radiotelemetric probe is reportedly as long as 4-7 days9, arterial pressure returns to normal much sooner, as demonstrated by the values reported here for blood pressure and heart rate. Indeed, these cardiovascular parameters are equivalent to those previously reported in similarly instrumented animals which were allowed up to 10 days to recover from surgery8,10.
The choice to use radiotelemetric probes for blood pressure measurement over a fluid-filled catheter was deliberate, as this reduces stress in the mice and also yields more clear and reliable blood and pulse pressure signals and heart rate values11. Using telemetric technology to record blood pressure poses an additional advantage since the need to frequently flush and maintain the fluid filled arterial catheter with heparinized saline, which inevitably disturbs the animal, is completely eliminated. Also, the approach of surgically exteriorizing, anchoring and protecting the intravenous catheter and bipolar electrode leads is ideal compared to other reports describing temporary storage of leads in a subcutaneous pocket12, since our approach avoids even brief re-anesthesia and surgical manipulation of the animal immediately before experimental recording, which would undoubtedly perturb the mouse and compromise the quality and interpretability of exquisitely sensitive autonomic nervous system data.
This method yields true RSNA signals, the quality of which are demonstrated by the characteristic bursts of electrical activity clearly distinguishable from background noise in a relaxed, quietly resting mouse. In addition, RSNA showed typical responsiveness to physical activity in the animal such as grooming and quiet eating as reported in the literature13,14. Given the characteristic increases in RSNA expected with natural movement or alertness of the animal, it is thus imperative to note and exclude these periods of time for the purpose of experimental analysis and to focus on segments of the recording during which the animal is quietly resting. This helps to prevent possible misinterpretation of the data. Other factors which can lead to data misinterpretation include electrical noise or interference, as well as signal contamination with ECG pulses15. Excessive movement of the exteriorized portion of the electrode leads can also influence the quality of the RSNA signal and can appear as an unstable or "wavering" baseline. At times these sources of signal interference can appear and spontaneously disappear during a perfectly clear recording and should be excluded from analysis5,15,16. An additional consideration is the time at which the recordings are obtained. It is important to note that blood pressure and RSNA do vary with the circadian rhythm, so it is ideal to conduct experiments at the same time of the day to avoid this potentially confounding factor. In this study, we did not observe significant variability of blood pressure and RSNA due to circadian oscillations as we recorded all parameters between 10 am and 6 pm – well within the daylight cycle of the animal housing facility. Another important component of this report is the validation of the RSNA signal, which as demonstrated, is indeed entrained with the arterial baroreflex. Given the rapid reduction and elevation of RSNA in parallel with the pharmacologically-induced drop and increase in systemic blood pressure, the arterial baroreflex was most certainly intact – which itself demonstrates that occlusive implantation of the radiotelemetry catheter in one carotid artery does not interfere with normal cardiovascular function. The virtual disappearance of the RSNA signal upon ganglionic blockade with hexamethonium further confirms recording of postganglionic RSNA.
It would be ideal to provide a longer post-surgery recovery period for the mice, however we and others in this field recognize that maintaining long-term viability of autonomic nerves in chronically instrumented animals, especially mice, remains challenging. Although the RSNA signal quality diminished over the course of several days post-surgery, it was still possible to reliably record true RSNA for at least 3 consecutive days in all mice and for up to 5 days in about half of the animals. This accomplishment in itself signifies a breakthrough in the field of autonomic studies in mice. Furthermore, this method maximizes the use of precious transgenic animals, as it is possible to record multiple experimental and control trials in the same animal on different days, of course, allowing for randomization of trial order and proper baseline recording before each experiment17. It is encouraging to see successful reports of long-term sympathetic nerve recordings conducted in conscious rodents18,19,20 including advancements in telemetric nerve recording technologies for rats15,21. Miniaturization of this technology for use in the conscious mouse is forthcoming and in the meantime, we strive to improve this technique to increase the longevity of the sympathetic nerve fibers to extend the experimental window and perhaps permit a longer post-surgery recovery time. However, this method will remain a useful and readily accessible and affordable alternative/complement to any future developments in telemetric nerve recording technology in mice, which do require an investment in dedicated equipment and regular device maintenance.
The need for reliable techniques for assessing cardiovascular and autonomic function in mice has never been so great, considering the ever-growing interest in transgenic mouse models in the field of biomedical research. Great strides have been made in many areas of physiology, however there is still far to go in terms of standardizing and optimizing approaches for evaluating autonomic function in the mouse. To date, there is one report describing measurement of sensory nerve activity in the conscious mouse12. This approach outlines the measurement of bladder sensory nerve activity and involves anesthesia and surgical manipulation of subcutaneously placed catheters immediately before experimental recording as well as physical restraint of the mice during the course of the experimental protocol12. These factors are known stressors that are completely avoided with the present approach, which can certainly be tailored for recording of a variety of nerves of interest in addition to renal nerves. More recently, sympathetic nerve measurements in conscious mice have been reported, however, these measurements are largely conducted hours following surgical preparation, with no mention of analgesic administration22. Aside from these reports, autonomic function has been otherwise assessed exclusively in anesthetized mice. A thorough review of the literature yields a multitude of approaches, hours-long experimental duration, anesthetic combinations/doses, mechanical ventilation and often creative measures taken to sustain the mice in a state bearing some semblance to the physiological (i.e. oxygen blown directly toward the animal's nose)23,24,25,26,27,28,29,30,31. Amongst these studies, reports of blood pressure values are absent, or abysmally low – below the physiological range of systemic arterial pressures2. This is problematic on many levels, but especially so when proper assessment of autonomic function in these animals is concerned, given the established link between blood pressure and autonomic tone. Anesthetic agents themselves directly impact sympathetic tone, with many reports suggesting that anesthesia dampens sympathetic activity. Indeed, evidence demonstrate that urethane, the most widely chosen anesthetic for acute nerve recording experiments32, dose dependently decreases RSNA33 and inhibits the arterial baroreflex34. Conversely, other reports suggest that urethane increases sympathetic tone35. Granted, such studies typically compare experimental nerve activity as a change from a recorded baseline, however the altered state of the autonomic nervous system under the above-described conditions undeniably precludes detection of discrete changes in nerve activity.
The challenge of this method lies mainly in the surgical skill required for successful preparation of the mouse for conscious nerve recording. However the investment in honing these skills is more than compensated by the quality and reliability of the direct RSNA data produced. This approach completely circumvents the limitations posed by indirect assessments of autonomic control such as plasma catecholamine levels, which are quite labile in mice and are limited by the amount of blood that may be humanely collected36. In addition, plasma catecholamine level as well as pharmacological autonomic blockade estimate overall autonomic tone1 as opposed to the discrete contributions of specific nerve populations, which are generally of more interest. Mathematic evaluation of autonomic tone via power spectral analysis of blood pressure and heart rate traces is useful for evaluating autonomic function in human subjects, however this technique may not be adaptable for mice36,37. Therefore, direct sampling of nerve activity in a conscious, comfortably resting mouse is ideal as it closely reflects the natural, intact autonomic status of the subject and facilitates sophisticated evaluation of the contribution of selected nerves to physiological phenomena of interest.
The authors have nothing to disclose.
S.M.H. was supported by postdoctoral fellowships from the Canadian Institutes for Health Research (CIHR), Heart & Stroke Foundation of Canada (HSFC) and Alberta Innovates Health Solutions (AiHS); J.E.H. is supported by a grant from the National Heart, Lung and Blood Institute PO1HL-51971.
Teflon-coated stainless steel multiple stranded wire | A-M Systems | 793200 | 0.001in diameter bare; 0.0055in diameter coated |
#11 Scalpel Blade | Fisher Scientific | ALMM9011 | |
Soldering Iron and solder | Any make or model suitable | ||
Male miniature pin connectors | A-M Systems | 520200 | Brass with gold plating |
Female miniature pin connectors | A-M Systems | 520100 | Brass with gold plating |
Heat Shrink tubing | Radio Shack | Model #: 278-1610 | Catalog #: 2781610 | 1.6 mm diameter |
Polyethylene 90 (PE90) tubing | VWR | CA-63018-703 | 0.86mm inner diameter; 1.27mm outer diameter |
Dissecting microscope | Leica Microsystems | Leica M80 | Any make or model also suitable |
Polyethylene 10 (PE10) tubing | Braintree Scientific | PE10 50 FT | 0.28mm inner diameter; 0.61mm outer diameter |
Super Glue Liquid | Loctite | n/a | Liquid Formula; any brand suitable |
Super Glue Gel | Loctite | n/a | Gel Formula; any brand suitable |
Polyethylene tubing | Scientific Commodities | BB31695-PE/13 | For pedestal 2.7mm inner diameter; 4.0mm outer diameter |
Hospital Sterilization Services & Ozone Sterilization packets | Contact local hospital sterilization services | ||
Isoflurane anesthesia | Abbott | 05260-05 | |
Deltaphase isothermal heat pads & surgical table | Braintree Scientific | 39OP | Keep heat pads warm in a 37°C water bath; Corresponding surgical table essential |
Glycopyrrolate | Amdipharm Mercury Company Limited | n/a | |
Isoflurane vaporizer system & flow gauge | Braintree Scientific | VP I | Include medical grade oxygen supply |
Tissue scissors | Fine Science Tools | 14173-12 | |
Fine spring scissors | Fine Science Tools | 15006-09 | |
Small cotton-tipped applicators | Fisher Scientific | 23400100 | |
Fine Straight Forceps | Fine Science Tools | 11254-20 | #5, FST by Dumont Biologie Tip |
Angled Forceps | Fine Science Tools | 11251-35 | #5/45 FST by Dumont |
Small Absorbent Spears | Fine Science Tools | 18105-03 | |
Parafilm | Sigma Aldrich | BR701605 ALDRICH | |
Kwik-Sil 2 component Silicone Polymer | World Precision Instruments (WPI) | KWIK-SIL | Purchase extra specialized tips from WPI |
5-0 Polysorb Suture | Tyco Healthcare | n/a | |
6-0 Silk Suture | Braintree Scientific | SUT-S 104 | Deknatel brand, spool |
Radiotelemetry Probe | Data Sciences International (DSI) | TA11-PAC10 | |
Radiotelemetry Receiver | Data Sciences International (DSI) | PhysioTel RPC-1 | |
Ambient Pressure Reference | Data Sciences International (DSI) | Apr-01 | |
Pressure Output Adapter | Data Sciences International (DSI) | R11CPA | |
Rena Pulse Tubing | Braintree Scientific | RPT-040 | |
Infusion Swivel | Instech Solomon | 375/D/22 | |
Swivel Support Arm & Mount | Instech Solomon | SMCLA | |
Polysulfone button | Instech Solomon | LW62S/6 | |
Stainless steel spring | Instech Solomon | PS62 | |
Vetbond surgical adhesive | 3M | n/a | |
Triple Antibiotic Ointment | Fougera | n/a | |
PowerLab 8 Channel Data Acquisition System & Software | ADInstruments | PowerLab 8/35 | |
PVC Insulated Cable | Belden | PVC Audio Connection Cable 32 AWG | |
Preamplification Headstage | Dagan Corporation | Model 4002 | |
Differential Amplifier | Dagan Corporation | EX4-400 | |
Sodium Nitroprusside | Sigma Aldrich | 71778-25G | |
Phenylephrine | Sigma Aldrich | P6126-5G | |
Sterile Physiological Saline 0.9% NaCl | Beckton Dickinson | Contact local hospital supplier | |
hexamethonium | Sigma Aldrich | H0879-5G | |
Stainless Steel top anti vibration table | n/a | n/a | Custom designed in-house; Solid steel plate on a benchtop is also suitable |
Faraday cage | n/a | n/a | Custom designed and constructed in-house |
Small animal hair trimmer | n/a | n/a | Drugstore, men's beard trimmer suitable |
Dipilatory Cream | n/a | n/a | Veet brand, sensitive skin formula |
10% Povidone Iodine | Purdue Products | Betadiene | |
70% Ethanol | n/a | n/a | |
Steel microretractors | n/a | n/a | Made in-house. Bend a steel paper clip & loop 4-0 silk to form a retractor |
Hemostats | Fine Science Tools | 13011-12 | |
Heat Gun | Fisher Scientific | 09-201-27 |