ILAR Journal Online, Volume 43(3) 2002: Advanced Physiological Monitoring in Rodents

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ILAR Journal V43(3) 2002
Advanced Physiological Monitoring in Rodents

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Miniaturization

Miniaturization: An Overview of Biotechnologies for Monitoring the Physiology and Pathophysiology of Rodent Animal Models
Tamara L. Goode and Hilton J. Klein

Tamara L. Goode, D.V.M., M.S., who is now with AstraZeneca Pharmaceuticals in Wilmington, Delaware, was a Research Veterinarian in the Department of Laboratory Animal Resources at Merck Research Laboratories, West Point, Pennsylvania, during preparation of this article. Hilton J. Klein, V.M.D., M.S., is Senior Director of Comparative Medicine in the Department of Laboratory Animal Resources at Merck.

Abstract

Recent advances in bioengineering technologies have made it possible to collect high-quality reproducible data quantitatively in a wide range of laboratory animal species, including rodents. Several of these technologies are incorporated into a plan called Miniaturization, which aims to design, develop, and maintain rodent animal models to study the pathophysiology and therapy of human diseases. Laser Doppler flowmetry, digital sonomicrometry, bioelectrical impedance, and microdialysis are some of the most widely used methods under the plan because they cause minimal pain and distress, reduce the number of animals used in biomedical research, and allow chronic, nonterminal assessment of physiological parameters in rodents. An overview of each of these technologies and their major applications in rodents used for biomedical research is provided.

Key Words: animal alternatives; bioelectrical impedance; digital sonomicrometry; laser Doppler flowmetry; microdialysis; miniaturization

Introduction

With the increased use of genetically engineered animals in biomedical research, mice and rats are becoming the animal models of choice in many physiological experiments. Due to their small size, however, the in vivo measurement of clinical, anatomical, and physiological parameters has been difficult. Recent technological advances and the miniaturization of biomedical devices have made it possible to acquire physiological data in mice and rats that formerly could be collected only in larger laboratory animal species, such as the dog or monkey.

In our laboratory, we have developed a novel plan called miniaturization, which uses one or more biotechnological advances to develop rodent animal models for biomedical research. In addition to reducing and refining the number of animals used in biomedical research, this strategy is a cost-effective method to generate high-quality, reproducible physiological data for pharmaceutical research programs. An overview of the most common biotechnologies used under the plan is provided in addition to their major applications in biomedical research. We emphasize the use of each technology for long-term, chronic monitoring of physiological parameters in rodents. In this issue of ILAR Journal, other authors provide extensive reviews of other technologies incorporated in the Miniaturization plan. The focus of this article is to cover technologies not reviewed by others, which include laser Doppler flowmetry (LDF¹), digital sonomicrometry, bioelectrical impedance, and microdialysis.

LDF

LDF is a method for determining blood flow through tissue capillaries, arterioles, and venules. This technique is commonly used to monitor the effect of environmental conditions, physical manipulations, and drug treatments on tissue perfusion. It has been used extensively in biomedical research, and its methods have been found to correlate with validated methods of blood flow measurement, such as the hydrogen clearance, [¹⁴C]iodoantipyrine, and radioactive microsphere methods (Goadsaby 1991; Kirkeby et al. 1995; Kramer et al. 1996; Skarphedinsson et al. 1998). Unlike ultrasonic transit-time methods of measuring blood flow, LDF has the advantage of allowing nonterminal evaluation of microvascular perfusion in real time.

Laser Doppler detects shifts in the frequency of laser light (Doppler shift) after it interacts with moving components of tissue such as red blood cells (Bonner et al. 1981). The flow of blood is measured by directing a laser beam to an area of tissue to be investigated. Upon contact with red blood cells in the target tissue, light waves are reflected and scattered, resulting in broadening of the light wave frequency, which a photodetector detects. The greater the net movement within tissue, the greater the broadening of the frequency of light detected and the greater the Doppler shift. The photodetector converts the broadened light frequencies into an electrical signal that is processed to provide an estimation of blood flow, termed flux. Flux is proportional to the average speed and number of red blood cells in the area of light penetration, generally 1 mm³ or smaller (Figure 1). It is not possible to determine absolute measurements of blood flow because the changes in wavelength that occur are unrelated to their direction of movement of red blood cells.

Figure 1 Basic operating principles of laser Doppler flowmetry. A laser beam is directed to an area of tissue. Upon contact with red blood cells in the target tissue, light waves are reflected and scattered, resulting in broadening of the light wave frequency, which is detected and received by a photodector.

The two types of laser Doppler instruments used to measure blood flow are single point fiber optic monitors and laser Doppler images (Figure 2). The single point fiber optic monitor is the instrumentation most frequently used to measure flux in rodents. Laser light is transmitted via a fiber optic probe placed in direct contact with the tissue under investigation. Several fiber optic probes are commercially available. Noninvasive continuous measurements of flux are made with skin surface probes. Needle probes are designed to penetrate tissues and therefore are more invasive than surface probes. They can be used intraoperatively to measure flow within tissues such as the brain or liver. Endoscopic probes are less invasive than needle probes and measure flow to mucosal surfaces.

Figure 2 Laser Doppler instrumentation. (A) Scanning laser Doppler imager; (B) single point fiber optic monitor with fiber optic probes.

The laser Doppler imager is a noninvasive system that measures flux within tissues without the use of fiber optic probes. The instrument scans tissues measuring 25 to 2500 cm² via a low-power collimated laser beam directed in a rectilinear pattern. A moving mirror directs reflected laser light from moving red blood cells to a photodetector. Subsequent processing results in a two-dimensional color-coded image of flux. The data in Figure 3 are representative of the data output seen with laser Doppler imaging in a mouse model of angiogenesis. Low flux level is expressed in blue, with green, yellow, and red indicating increasingly higher level regions of flux. Gray areas represent regions where no flux can be detected. Imaging is facilitated by scanning the subject against a black background for greater contrast and visualization of the flux color spectrum and to enhance image analysis. In experiments where repetitive analysis of blood flow is required in the same animal, the region of interest in which flux must be determined should be permanently marked to allow appropriate realignment and to ensure that the same region is scanned.

Figure 3 Laser Doppler images of a dorsal air sac mouse model of angiogenesis. (A) Photo and flux images of the dorsal surface of a control animal 4 days after implantation of a diffusion chamber containing phosphate-buffered saline. Low levels of flux were detected around regions where the diffusion chamber was implanted. (B) Photo and flux images of a mouse 4 days after implantation of a diffusion chamber containing tumor cells known to promote angiogenesis. Note the significant increases in flux, as depicted by the increased pixels of red that were observed around the region where the diffusion chamber was implanted. (Images obtained using Moor Instruments Laser Doppler Imager, Moor Instruments, Wilmington, Delaware.)

Laser Doppler requires the tissue measured to remain stationary in relation to the laser beam. Movement of the animal can cause displacement of the laser Doppler flow probe during measurement and a subsequent shift of the baseline blood flow measurement (Dirnagl et al. 1989). Stability and accurate reaffixation of the probe is critical to reduce variability. For this reason, anesthesia of the animal is often required. Various anesthetics have been shown to alter blood flow in rodents (Dalkara et al. 1995; Kiatchoosakun et al. 2000; Koorn et al. 1993; Lindauer et al. 1993; Okamota et al. 1997; Short 1987). Therefore, the anesthetic regimen used during imaging should be tailored to the experimental protocol and should not adversely affect the vascular bed to be examined. Perfusion is temperature dependent, and fluctuations in body temperature may affect anesthetic requirements and exert confounding vasoactive properties (Kuluz et al. 1993). Body temperature should be monitored frequently and regulated via the use of an isothermic heating pad.

LDF has been used for ocular, cerebral, cutaneous, auricular, splanchnic, and renal blood flow in a wide range of laboratory animal species. A list of laser Doppler applications is provided in Table 1. With respect to rodents, the technology has been described in the mouse, rat, hamster, gerbil, guinea pig, and chinchilla.

The measurement of regional cerebral blood flow in the rat is the most widely reported experimental use of LDF. The single point laser Doppler is the instrument most frequently used for cerebral flow measurements; however, methods using the laser Doppler imager have been described (Kimme et al. 1997). When using the single point laser Doppler device, needle probes are typically advanced through a craniotomy to the cerebral cortex at a depth of approximately 0.5 mm and a measuring hemisphere of 0.5 to 1.0 mm (Barfod et al. 1997; Fabricius et al. 1999). Although most measurements using single point laser Doppler are achieved by advancing a flow probe through a craniotomy, reproducible and reliable measurements have also been described using a partial craniotomy in which a thin layer of calvarium is left intact (Gu et al. 1999). Similar cerebral flow measurements have also been described in the mouse (Gong et al. 1998; Nawashiro et al. 2000). Unlike the rat, surface measurements of cerebral flow can also be made in the mouse through the intact skull bone (Kamii et al. 1994).

Quantification of changes in regional cerebral blood flow with laser Doppler is commonly performed in the rat, although utility has also been documented in the mouse, hamster, gerbil, and guinea pig (Beattie et al. 1993; Connolly et al. 1996; Czernicki et al. 1996; Dalkara et al. 1995; Duan et al. 1994; Huang et al. 1999; Ichihara et al. 1996; Kilic et al. 2000; Maeda et al. 1999). In these models, cerebral blood flow measurements are typically performed over short periods in anesthetized animals because long-term monitoring is associated with increased mortality due to long anesthetic durations (Gu et al. 1999). Noteworthy are the applications of laser Doppler flowmetry in which measurement of cerebral blood flow was performed in unanesthetized rats (Gu et al 1999; Sato et al 1994). A metal guide cannula with customized flow probe was permanently anchored to the skull for continuous, repetitive measurements of cerebral blood flow. Upon recovery from surgery, probes were advanced through the adapter to measure flow. Probes were removed and repositioned at exactly the same anatomical location for repeated measurements in the cortex for up to 4 days.

The second most common application of LDF is in the area of angiogenesis. Angiogenesis plays a critical role in the growth, maturation, and maintenance of normal and neoplastic tissues (Moses and Langer 1991). Disease processes in which angiogenesis can lead to pathology are tumor development, rheumatoid arthritis, psoriasis, and diabetic retinopathy (Folkman 1995; Moses and Langer 1991). Pharmacological agents that prevent angiogenesis (antiangiogenesis) are promising tools to restrict tumor growth, prevent metastasis, and treat disease. The laser Doppler imager is used widely in several animal models to characterize angiogenic responses after therapeutic interventions. The most common models in which this technique is applied include the hamster cheek pouch (Colantuoni and Bertuglia 1997; Orlandi et al. 1986), disc angiogenesis (Fajardo et al. 1988; Kowalski et al. 1992), rat and mouse dorsal air sac (Lichtenberg et al. 1997), matrigel (Passaniti et al. 1992), and mouse and rat hind limb ischemia (Couffinhal et al. 1998; Lloyd et al. 2001; Skjeldal et al. 1993; Taniyama et al. 2001). A chorioallantoic membrane model using fertilized eggs has been described with laser Doppler imaging; however, it has been considered a less relevant model to predict the human physiology and pathology of angiogenesis because it is not a mammalian system (Kowalski et al. 1992). Of all the models described, the mouse and rat hind limb ischemia models are the most commonly studied. Postischemic and reperfusion microcirculatory changes are evaluated after unilateral femoral artery occlusion (Lloyd et al. 2001; Taniyama et al. 2001). The reduction in hind limb perfusion followed by subsequent collateral vessel development is often monitored (Couffinhal et al. 1998; Kanno et al. 1999; Lloyd et al. 2001; Skjeldal et al. 1993; Takeshita et al. 1998; Taniyama et al. 2001).

LDF is a valuable tool for real-time assessment of microcirculation. Combining laser Doppler with other biotechnologies may achieve increased value. Kaiser and During (1995) combined laser Doppler flowmetry with microdialysis as a novel approach to evaluate regional cerebral flow in correlation with neuronal activity, allowing greater amounts of physiological data to be generated in fewer numbers of animals.

Digital Sonomicrometry

Digital sonomicrometry is an ultrasonic measurement system that utilizes piezoelectric crystals to measure multiple distances within an aqueous medium or soft tissue. Distance measurements are determined by implanting crystals that transmit and receive sound energy at ultrasonic frequencies of greater than or equal to 1 MHz (Espisito et al. 2000; Gorman et al. 1996; Hansen et al. 1995).

Digital sonomicrometry has been used extensively for nonterminal assessment of cardiovascular function in large animal species such as the dog. Similar parameters can be monitored in rodents due to the miniaturization of transducers used in the digital sonomicrometry system. Applications for digital sonomicrometry are provided in Table 2.

The three major components in a digital sonomicrometry system include the piezoelectric crystals, the sonomicrometer, and the computer acquisition system. Piezoelectric crystals are cylindrical in shape, attached to Teflon-coated copper or stainless steel wire, and encapsulated in an epoxy resin (Figure 4). Crystals, which range in size from 0.5 to 2.0 mm, can be implanted in animals as small as mice. Multiple distances can be measured at 200 times per second and with 0.006-µm resolution. Crystals are connected to a sonomicrometer, which is used in conjunction with a customized computer acquisition system to measure distances in tissues.

Figure 4 Digital sonomicrometry instrumentation. (A) (Top) bare and (bottom) epoxy coated piezoelectric crystals. Crystals are cylindrical in shape, attached to Teflon-coated copper or stainless steel wire, and encapsulated in epoxy resin. Crystals used in rodents typically range in size from 0.5 to1.0 mm. (B) Four-channel digital sonomicrometer with two piezoelectric crystals.

Distances are measured by implanting a network of two or more piezoelectric crystals. Crystals are connected to the sonomicrometer, and an electrical impulse is generated resulting in oscillation and production of ultrasonic signal by one of the crystals in the network. This signal is transmitted to additional crystals, which act as piezoelectric receivers. The time from when the sound is broadcast by the active transmitter to the instant it was detected by each corresponding receiver is determined. The velocity of ultrasound through the tissue of interest is known, and the exact distance measurement between crystals is calculated by the system using the formula distance is equal to velocity multiplied by time (d = v × t).

Crystals in the digital sonomicrometry system are unique in that they alternate between transmitting and receiving modes (Figure 5). This characteristic allows multiple distances to be measured in the tissue of interest without crystal alignment and sonomicrometer calibration, which saves substantial time on experimental setup. Data acquired with digital sonomicrometry crystals grow exponentially with the number of crystals used (Figure 6). Conventional analog sonomicrometry systems differ and require implanting the piezoelectric crystals in pairs because one crystal serves as a transmitter and the other as a receiver. Crystals can only receive and transmit signals within the predetermined pair, and only distances within the paired group can be measured. Data collection requires extensive labor-intensive calibration using oscilloscopes before implantation of the crystal pairs.

Figure 5 Basic operating principles of digital sonomicrometry. A network of two or more piezoelectric crystals is implanted onto a tissue of interest. Electrical impulses are directed to a crystal resulting in oscillation and production of an ultrasonic signal, which is transmitted to other additional crystals in the network. Crystals alternate between transmitting and receiving modes. Left (t₁): 1 = transmitting crystal, 2 and 3 = receiving crystals; center (t₂): 2 = transmitting crystal, 1 and 3 = receiving crystals; right (t₂): 3 = transmitting crystal, 1 and 2 = receiving crystals.

Figure 6 Exponential growth of crystals. Data acquired with the digital sonomicrometry system grow exponentially with the number of crystals implanted.

Sonomicrometry has been used in a variety of laboratory animal species to determine cardiac, vascular, pulmonary, gastrointestinal, urogenital, and musculoskeletal measurements. Most investigations have been described in birds, mice, rats, dogs, pigs, and nonhuman primates. Applications in rodents are currently limited to rats and mice in cardiovascular research. Left ventricular dimension and function are typically evaluated through the use of isolated heart preparations and in acute studies in anesthetized animals.

The end systolic pressure-volume relation (ESPVR¹) has been the standard method for evaluating cardiac function in large animals (Kubota et al. 1997). Contractile function can also be assessed in rodents by establishing pressure-dimension relations in vivo. In the absence of regional wall motion abnormalities, dimension can be substituted for volume (Espisito et al. 2000; Kubota et al. 1997).

Determination of ESPVR in rodents has been well characterized in genetically engineered rodents (Espisito et al. 2000; Hajjar et al. 1998; Hoit and Walsh 1997; Kubota et al. 1997; Miyamoto et al. 1999; Schmidt et al. 2000). To measure ESPVR, animals must be anesthetized, intubated, and mechanically ventilated. Left ventricular pressure measurements are made with a micromanometer, which is placed directly into the apex of the heart or advanced to the ventricle through the carotid artery (Hoit 2001). Via a thoracotomy incision, two to four crystals are affixed to the left ventricle to measure dimension (Hajjar et al. 1998; Kubota et al. 1997). Typically two crystals are placed on the anterior and posterior walls of the left ventricle to determine short-axis dimension, with two crystals at the base and apex to measure long-axis dimension.

Espisito and colleagues (2000) have described a new technique to measure contractility in the mouse. The procedure involves placement of orthogonal crystals on the endocardial surface of the heart and placement of two additional crystals on the base and apex of the epicardial surface of the left ventricle. Using this method, internal dimensions of the ventricle can be obtained in two planes, allowing absolute left ventricular volume to be calculated. Cardiac function has also been determined in rodents via conductance volumetry and echocardiography methods. The sonomicrometry technique is comparable to these systems (Espisito et al. 2000). Excellent spatial and temporal resolution is possible with the use of sonomicrometry, and placement of an orthogonal crystal on the endocardial surface juxtaposed to a crystal on the epicardial surface can be performed for instantaneous measurements of wall thickness and calculation of wall stress.

Methods for examining cardiac function in unanesthetized rodents are needed. In larger animals, the wires connected to crystals are tunneled subcutaneously and later attached to a tether system for instantaneous measurements of left ventricular cavity dimension. Recently, miniaturized piezoelectric crystals have been modified to include suture material within the Teflon coating. This modification allows permanent fixation of crystals to the epicardial surface of the heart and theoretically makes chronic measurement of left ventricular function in unanesthetized rodents feasible. Methods for evaluating left ventricular pressure chronically in unanesthetized rodents still must be developed.

Digital sonomicrometry has limitations. Instrumentation requires thoracotomy, which is technically challenging and requires great skill in microsurgical technique. Sonomicrometry is insensitive to minute focal wall motion abnormalities such as infarcts and therefore may not be a reliable methodology for study of cardiac ischemia and reperfusion. Although sonomicrometry is very invasive, the resolution is improved compared with other conventional methods. The technique will continue to offer value in characterizing cardiovascular phenotypes in genetically engineered rodents.

Bioelectrical Impedance

Bioelectrical impedance analysis is a rapid noninvasive technique for estimating fluid distribution and body composition (Kushner and Schoeller 1986). Although extensively validated in humans, the application of bioelectrical impedance in biomedical research has been limited to rats, cats, dogs, sheep, and nonhuman primates (Hoffer et al. 1969; Ilagan et al. 1993; Jenkins et al. 1988; Kichul et al. 1995; Lukaski et al. 1986; Mathers et al. 2000; Olde Rikkert et al. 1997; Scheltinga et al. 1991; Stanton et al. 1992). Measurements are made using and impedance analyzer similar to the one in Figure 7.

Figure 7 A multifrequency bioimpedance analyzer used to evaluate fluid distribution and body composition in laboratory animals (Xitron Technologies, San Diego, California).

Bioelectrical impedance assumes that the complex geometry of the living body can be approximated as a simple cylindrical conductor in which low-voltage electrical currents are delivered at a single fixed frequency or multiple frequencies. It assumes that resistance or impedance to these currents is inversely proportional to the amount of total body water and lean body mass present. Aqueous tissues (fat-free mass) such as lean muscle and fluids are considered excellent conductors, whereas fat and bone have poor conductance properties.

Water is the most abundant component of the body and accounts for 60% of its weight (Cornish et al. 1993). Total body water, which is located primarily in fat-free mass, is divided into extracellular fluid (ECF¹) and intracellular fluid (ICF¹) compartments. In determining fluid distribution and body composition, bioelectrical impedance assumes that ICF and ECF behave as conductors and that cell membranes and tissue interfaces act as imperfect capacitors. Single-frequency bioelectrical impedance delivers a constant alternating current at a fixed frequency typically measuring 50 kHz. A derivative of this method is multifrequency bioelectrical impedance analysis (MFBIA¹), which delivers alternating currents at frequencies ranging from 1 kHz to 1 MHz. Low- and high-frequency currents in MFBIA help distinguish ECF and total body water (TBW¹) because at low frequencies (<1kHz), current passes mainly through the ECF, and at higher frequencies (100 kHz to 1 MHz), the current penetrates both ECF and ICF spaces (predicting TBW). Total body impedance is related to the amount of water in the body. When alternating current passes through the body, the fluid is the main conductive pathway. By determining the length of the conductor, volume of total body water can be predicted in accordance with methods established by Cornish and coworkers (1992) in which algorithms are generated to predict total body water and fat-free mass in several populations of animals.

MFBIA is the most frequently used instrument in biomedical research. Measurements are made in anesthetized animals placed on a nonconductive surface and with electrodes inserted subcutaneously in a tetrapolar arrangement. Electrodes can be fabricated from stainless steel hypodermic needles, or platinum transdermal electrodes can be used. Two source electrodes and two detecting electrodes are placed subcutaneously along the midline, with the source electrodes placed dorsally at the back of the head and base of the tail and the detecting electrodes placed ventrally over the xiphoid and pelvic bone. Interelectrode length (length of the conductor) of the animal is determined for future TBW analysis.

ICF and ECF compartments are tightly regulated unless trauma occurs or pathological conditions such as pulmonary disease, cardiovascular disease, renal disease, obesity, immunodeficiency, and sepsis develop (Suzuki and Wilmore 1993). Management of dysregulation of fluid compartments becomes important in achieving homeostasis and identifying and treating fluid imbalance. Bioelectrical impedance analysis becomes a particularly useful tool in biomedical research for assessing changes in fluid distribution. Increases in ECF volumes have been determined with MFBIA after injection of aliquots of saline in the intraperitoneal cavity (Cornish et al. 1994). Further studies are needed to evaluate the ability of MFBIA to detect fluid shifts after therapeutic compounds are administered.

Several methods have been described to measure body composition (Table 3). A potential advantage of impedance analysis over these methods is its ability to measure the distribution of water between ICF and ECF spaces. The validity of MFBIA as a tool in measuring body composition has been established in the rat. Suzuki and Wilmore (1993) determined a correlation between resistance signals of MFBIA and body composition.

MFBIA is an excellent tool for estimating changes in ECF and ICF. The technology proves to be a safe, rapid, reproducible technique with exceptional promise in biomedical research because it has been validated against traditional measures of body composition and fluid analysis. A great advantage of MFBIA is its low cost compared with other conventional methods of measuring body composition and fluid distribution.

Microdialysis

Microdialysis is an analytical technique used to evaluate changes in the chemical composition of tissues (de la Pena et al. 2000). It has been used extensively in pharmacokinetic and pharmacodynamic studies to measure unbound fluid concentrations in all tissues and organs of the body (Verbeeck 2000). Microdialysis does not require the removal of body fluids, therefore, exogenous and endogenous compounds present in the ECF can be evaluated without altering fluid balance or disturbing blood homeostasis (de la Pena et al. 2000).

The main equipment used for microdialysis is illustrated in Figure 8 and consists of a microdialysis probe with inlet and outlet tubing, infusion pump, animal housing chamber, fraction collector, analytical measuring device, and computer system. An array of microdialysis probes can be used with this technique. Flexible probes are used for sampling blood and soft peripheral tissue (i.e., muscle, skin, liver, tumors) and must be secured for uncompromised stable monitoring. Linear probes are used to measure neurotransmitter levels in the brain (Davies et al. 2000). Intracerebral probes are rigid linear probes that are typically advanced through a cannula surgically placed at least 24 hr before microdialysis sampling. Shunt probes, which are linear dialysis probes housed in plastic tubing, are used for continuous monitoring of moving fluids such as bile. These probes are normally implanted in rats and, unlike conventional methods of bile sampling, conserve bile salts during pharmacokinetic studies (Davies et al. 2000).

Figure 8 Equipment used for microdialysis procedures (fraction collector, animal holding chamber with tether system, infusion pump).

Probes consist of a semipermable membrane in the formation of a hollow fiber membrane. Tubing is affixed to an inlet and outlet port of the probe, and an aqueous solution (perfusate) similar to the composition of tissue sampled is infused via the infusion pump. During sampling, animals are placed into a tethering system so that the animal can move about freely without tangling the microdialysis tubing. A fraction collector is used to collect microdialysis samples. The volume and time samples collected by the fraction collector are predetermined and controlled by a computer system. Alternatively, samples can be delivered directly to an analytical measuring device, in which case an online injector is used. Devices that can accommodate small volumes and detect low concentrations of microdialysis samples are used for analysis. Examples of analytical devices include high-pressure liquid chromatography and mass spectroscopy.

Microdialysis sampling is performed by surgically implanting a probe into an organ or biological fluid of interest. Perfusate similar to the ionic composition and pH of the medium sampled is infused through the interior of the probe membrane. The driving force for moving substances across the microdialysis probe membrane relies on a concentration gradient generated between the ECF space and the probe lumen. Substances with a molecular weight less than the molecular weight of the microdialysis membrane will diffuse across the membrane whereas molecules with molecular weights greater than that of the membrane are excluded from the perfusate. Once a molecule diffuses across the probe membrane, it is protected from further metabolism by enzymes in the tissue. Such protection is not possible using blood sampling or open cannula methods. The solution that diffuses out of the probe is representative of the concentration of the ECF and is termed the dialysate. The dialysate is collected into a fraction collector or can be directly transported to an analytical system. Analytes that are typically measured with microdialysis include drugs and their metabolites, neurotransmitters, and amino acids.

Microdialysis procedures are not performed under equilibrium, and the concentrations of drugs in the dialysate collected will differ from concentrations around the probe (Davies 1999). Due to this partial recovery of analytes, tedious in vitro calibration methods are required to determine absolute concentrations of substances in the ECF. It is important to consider several factors when performing microdialysis procedures. These factors include probe type and composition, perfusate flow rate and temperature, surgical recovery time, and length and composition of tubing.

Probe materials vary in size and composition, and the type used in an investigation is dependent on the accessibility and nature of the target tissue to be sampled (Davies et al. 2000). Materials used for probe membranes include cellulose, copolymers, acrylic copolymers, and polysulfone. It is important to choose an appropriate membrane or material that does not interact with the drug and does not adversely affect the concentration of the dialysate. Lipophilic drugs have a tendency to adhere to dialysis probes and connecting tubing (de Lange et al. 2000). The length and inner diameter of the outlet tubing are important and should be kept to a minimum to prevent a buildup of hydrostatic pressure across the probe membrane. It is recommended that inlet tubing to the probe have an inner diameter smaller than the probe itself and the outlet tubing from the probe have an inner diameter large than that of the probe (de Lange et al. 2000).

The composition and temperature of the perfusate used in microdialysis procedures are important and should mimic the tissue being sampled because alterations can occur in the dialysate (de Lange et al. 2000; Moghaddam and Bunney 1989). Most microdialysis procedures are performed with the perfusate at room temperature, which creates temperature gradients across the probe (de Lange et al. 2000). Methods to equilibrate the perfusate to body temperature have been used and should be considered (de Lange 1994). Flow rate of the perfusate will affect recovery. It is advantageous to perfuse at very low flow rates to obtain the highest concentration of analytes in the dialysate (de Lange et al. 2000). Flow rates of 0.1 to 1.0 µL/min are recommended (Davies et al. 1999). Typically probes are perfused at 1 µL /min, and 5 µL of the analyte is evaluated every 5 min.

Implantation of dialysis probes has been reported to elicit local inflammatory responses and minor physiological changes, therefore, it is best to give animals adequate time to recover before dialysate is collected. Recovery periods as short as 2 hr have been recommended, depending on the tissue sampled (de Lange et al. 2000).

Intracerebral microdialysis is the most frequently reported technique described in the literature. However, the technique has quickly expanded to include applications in pharmacokinetic and toxicology studies, in which other tissues like the skin, kidney, liver, and tumors are evaluated.

Neurotransmitter release is often evaluated in mice and rats via intracerebral microdialysis (Boschi and Scherrmann 2000). Procedures require stereotaxic surgery in which cannulae are implanted into a predetermined area in the brain. For chronic measurements, cannulae are secured with an acrylic resin and animals are allowed to recover a minimum of 24 hr before perfusate is infused. More than one microdialysis probe can be implanted into an animal, as in the case of work by Evrard and colleagues (1996), in which simultaneous blood and brain microdialysis probes were implanted to examine the effects of colchicine across the blood-brain barrier.

Simultaneous measurements have also been made with microdialysis probes in the blood along with multiple sights in the liver for investigations of phenol metabolism (Davies and Lunte 1996). In characterization of a gliosarcoma model, two intracranial microdialysis probes were implanted to evaluate tumoral brain metabolism in freely moving rats. Authors showed significant increases in tumor pyruvate and lactate compared with normal brain tissue (Darbin 2000).

Microdialysis offers numerous benefits in biomedical research. The number of animals required in pharmacokinetic studies can be dramatically reduced through the use of this technique. Because homeostasis is not affected by the application of microdialysis, numerous samples can be taken from the same animal, thereby decreasing the number of animals used in biomedical research. In addition, continuous and repeated sampling can be performed in conscious, freely moving animals, providing the target tissue remains unchanged. The ability to perform pharmacokinetic studies in mice via blood microdialysis is a major benefit inasmuch as the technique is not associated with fluid loss. Microdialysis is the most affordable technique to monitor ECF spaces in unanesthetized animals.

Conclusion

Miniaturization and refinement of conventional techniques have made it possible to collect data in rodents that are comparable in quality and quantity to data attained in large animal species. When utilizing any physiological assay, accuracy and reproducibility of results are critical, and variables that affect physiological monitoring must always be considered.

All of the technologies described in this review have shown measurable benefit while studying disease processes and measuring drug responses. The physiological methods chosen must be tailored to the hypothesis in question, and often a combination of techniques will enhance the experimental design as well as the power of the experiment.

Each technology serves as a valuable alternative by allowing the number of rodents used in biomedical research to be reduced while causing minimal pain and distress and by replacing methods that required necropsy as an endpoint. Although it is reasonable to state that laboratory rodents will not replace nonrodent species in biomedical research, the use of these technologies in mice and rats can serve as a valuable adjunct in studying the mechanism and treatment of disease processes before conducting investigations in larger animal species. Finally, all methods provide a cost-effective means of measuring physiological parameters in rodents.

¹Abbreviations used in this article: ECF, extracellular fluid; ESPVR, end systolic pressure-volume relation; ICF, intracellular fluid; LDF, laser Doppler flowmetry; MFBIA, multifrequency bioelectrical impedance analysis; TBW, total body water.

References

Barfod C, Akgoren N, Fabricius M, Dirnagl U, Lauritzen J. 1997. Laser-Doppler measurements of concentration and velocity of moving blood cells in rat cerebral circulation. Acta Physiol Scand 160:123-132.

Beattie DT, McNeil DK, Connor HE. 1993. The influence of neurokinins and calcitonin gene-related peptide on cerebral blood flow in anesthetized guinea pigs. Neuropeptides 24:343-349.

Bonner RF, Clem TR, Bowen PD, Bowman RL. 1981. Laser Doppler continuous real-time monitor of pulsatile and mean blood flow in tissue microcirculation. In: Chen SH, Chu B, Nossal R, eds. Scattering Techniques, Applied to Supra-Molecular and Nonequilibrium Systems. New York: Plenum. p 685.

Boschi G, Scherrmann J. 2000. Microdialysis in mice for drug delivery research. Adv Drug Deliv Rev 45:271-281.

Colantuoni A, Bertuglia S. 1997. Correlation between laser Doppler perfusion monitoring and hematocrit in hamster cheek pouch microcirculation. Int J Microcirc Clin Exp 17:33-40.

Connolly ES Jr, Winfree CJ, Springer TA, Naka Y, Liao H, Shi Du Yan, Stern DM, Solomon RA, Gutierrez Ramos JC, Pinsky DJ. 1996. Cerebral protection in homozygous null ICAM-1 mice after middle cerebral artery occlusion: Role of neutrophil adhesion in the pathogenesis of stroke. J Clin Invest 97:209-216.

Cornish BH, Thomas BJ, Ward LC. 1993. Improved prediction of extracellular and total body water using impedance loci generated by multiple frequency bioelectrical impedance analysis. Physiol Med Biol 38:337-346.

Cornish BH, Ward LC, Thomas BJ. 1994. Alteration of the extracellular and total body water volumes measured by multiple frequency bioelectrical impedance analysis (MFBIA). Nutr Res 14:717-727.

Cornish BH, Ward LC, Thomas BJ. 1992. Measurement of extracellular and total body water of rats using multiple frequency bioelectrical impedance analysis. Nutr Res 12:657-666.

Couffinhal T, Silver M, Zheng LP, Kearney M, Witzenbichler B, Isner JM. 1998. Mouse model of angiogenesis. Am J Pathol 152:1667-1679.

Czernicki Z, Suzuki R, Nakagawa K, Hirakawa K, Endo S. 1996. Acetazolamide produced blood flow velocity changes measured by laser Doppler in gerbils with reduced CBF. Acta Neurochir 138:81-83.

Dalkara T, Irikura K, Huang Z, Panahian N, Mosskowitz MA. 1995. Cerebrovascular responses under controlled and monitored physiological conditions in the anesthetized mouse. J Cereb Blood Flow Metab 15:631-638.

Darbin O, Lonjon M, Quentien MH, Michiels JF, Grellier P, Negrin J, Rostain JC, Risso JJ. 2000. In vivo study of tumor metabolism: An application of new multi-probe microdialysis system in the striatum of freely moving rats grafted with C6 cells. Brain Res 88:121-127.

Davies M, Cooper JD, Desmond SS, Lunte CE, Lunte SM. 2000. Analytical considerations for microdialysis sampling. Adv Drug Deliv Rev 45:169-188.

Davies M, Lunte CE. 1996. Simultaneous microdialysis sampling from multiple sites in the liver for the study of phenol metabolism. Life Sci 59:1001-1003.

Davies MI. 1999. A review of microdialysis sampling for pharmacokinetic applications. Anal Chim Acta 379:227-249.

de la Pena A, Liu P, Derendorf H. 2000. Microdialysis in peripheral tissues. Adv Drug Deliv Rev 45:189-216.

De Lange ECM, Danhof M, De Boer AG, Breimer DD. 1994. Critical factors of intracerebral microdialysis as a technique to determine the pharmacokinetics of drugs in rat brain. Brain Res 666:1-8.

De Lange ECM, De Boer AG, Breimer DD. 2000. Methodological issues in microdialysis sampling for pharmacokinetic studies. Adv Drug Deliv Rev 45:125-148.

Dirnagl U, Kaplan B, Jacewicz M, Pulsinelli W. 1989. Continuous measurement of cerebral cortical blood flow by laser-Doppler flowmetry in a rat stroke model. J Cereb Blood Flow Metab 9:589-596.

Duan CG, Li HW, Xu LN, Xiu RJ. 1994. Influences of puerarin on microcirculatory disturbance in brain of hamsters. Asia Pac J Pharmacol 9:93-95.

Espisito G, Santana LF, Dilly K, Cruz JD, Mao L, Lederer WJ, Rockman HA. 2000. Cellular and functional defects in a mouse model of heart failure. Am J Physiol Heart Circ Physiol 279:H3101-H3112.

Evrard PE, Deridder G, Verbeeck RK. 1996. Intravenous microdialysis in the mouse and rat: Development and pharmacokinetic application of a new probe. Pharm Res 13:12-17.

Fabricius M, Akgore N, Dirnagl U, Lauritzen M. 1999. Laminar analysis of cerebral blood flow in cortex of rats by laser-Doppler flowmetry: A pilot study. J Cerebr Blood Flow Metab 17:1326-1336.

Fajardo AF, Kowalski J, Kwan HH, Prionas SD. Allison AC. 1988. The disk angiogenesis system. Lab Invest 58:718-724.

Folkman J. 1995. Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat Med 1:27-31.

Goadsaby PJ. 1991. Characteristics of facial nerve-elicited cerebral vasodilatation determined using laser Doppler flowmetry. Am J Physiol 260:R255-R262.

Gong C, Qin Z, Betz AL, Liu XH, Yang GY. 1998. Cellular localization of tumor necrosis factor alpha following focal cerebral ischemia in mice. Brain Res 801:1-8.

Gorman JH, Gupta KB, Steicher JT, Gorman RC, Jackson BM, Ratcliffe MB, Bogen DK, Edmunds LH. 1996. Dynamic three-dimensional imaging of the mitral valve and left ventricle by rapid sonomicrometry array localization. J Thorac Cardiol Surg 112:712-725.

Gu W, Jiang W, Brannstrom T, Wester P. 1999. Long-term cortical CBF recording by laser-Doppler flowmetry I awake freely moving rats subjected to reversible photothrombotic stroke. J Neurosci Methods 90:23-32.

Hajjar RJ, Schmidt U, Matsui T, Guerrero JL, Lee K, Gwathmey JK, Dec G, Semigran MJ, Rosenzweig A. 1998. Modulation of ventricular function through gene transfer in vivo. Proc Natl Acad Sci U S A 95:5251-5256.

Hansen B, Menkis AH, Vesley I. 1995. Longitudinal and radial distensibility of the porcine aortic root. Ann Thorac Surg 60:S384-S390.

Hoffer EC, Meador CK, Simpson DC. 1969. Correlation of whole body impedance with total body water volume. J Appl Physiol 27:531-534.

Hoit B. 2001. New approaches to phenotypic analysis in adult mice. J Mol Cell Cardiol 33:27-35.

Hoit BD, Walsh RA. 1997. In vivo echocardiographic assessment of left ventricular function in transgenic and gene-targeted cice. Trends Cardiovasc Med 7:129-134.

Huang J, Kim LJ, Poisik A, Pinsky DJ, Connolly ES Jr, Muizelaar JP, Mayberg MR, Dacey RG Jr, Findlay JM. 1999. Titration of postischemic cerebral hypoperfusion by variation of ischemic severity in a murine model of stroke. Neurosurgery 45:328-333.

Ichihara SI, Tsuda Y, Hosomi N, Kitadai M, Matsuo H. 1996. Nimodipine improved brain energy metabolism and blood rheology during ischemia and reperfusion in the gerbil brain. J Neur Sci 144:84-90.

Ilagan J, Bhutani V, Archer P, Lin P, Jen KL. 1993. Estimation of body composition changes during weight cycling by bioelectrical impedance analysis in rats. J Appl Physiol 74:2092-2098.

Jenkins TG, Leymaster KA, Turlington LM. 1988. Estimation of fat-free soft tissue in lamb carcasses by use of carcass and resistive impedance measurements. J Anim Sci 66:2174-2179.

Kaiser MG, During MJ. 1995. Combining laser Doppler flowmetry with microdialysis: A novel approach to investigate the coupling of regional cerebral blood flow to neuronal activity. J Neurosci Methods 60:165-173.

Kamii H, Kinouchi H, Sharp FR, Koistiaho J, Epstein CJ, Chan PH. 1994. Prolonged expression of hsp70 mRNA following transient focal cerebral ischemia in transgenic mice over-expressing CuZn-superoxide dismutase. J Cerebr Blood Flow Metab 14:478-486.

Kanno, S, Nobuyuki O, Mayumi A, Saito S, Hori, K, Handa Y, Tabayashi K, Sato Y. 1999. Establishment of a simple and practical procedure applicable to therapeutic angiogenesis. Circulation 99:2682-2687.

Kiatchoosakun S, Kirkpatrick D, Walsh R, Hoit B. 2000. The effect of avertin anesthesia on the echocardiographic assessment of left ventricular function. J Cardiol Fail 6(Suppl 2):13.

Kichul C, Hill AG, Rounds JD, Wilore DW. 1995. Multifrequency bioelectrical impedance fails to quantify sequestration of abdominal fluid. J Appl Physiol 78:736-739.

Kilic E, Hermann DM, Hossmann KA. 2000. Recombinant tissue-plasminogen activator-induced thrombolysis after cerebral thromboembolism in mice. Acta Neur 99:219-222.

Kimme P, Gustafsson U, Sollevi A, Nilsson G, Sjoberg F. 1997. Cerebral blood flow of the exposed brain surface measured by laser Doppler perfusion imaging. Acta Physiol Scand 159:15-22.

Kirkeby OJ, Rise IR, Nordsletten L, Skjeldal S, Hall C, Risoe C. 1995. Cerebral blood flow measured with intracerebral laser-Doppler-flow probes and radioactive microspheres. J Appl Physiol 79:1479-1486.

Koorn R, Kahn RA, Brannan TS, Martinez Tica J, Weinberger J, Reich DL. 1993. Effect of isoflurane and halothane on in vivo ischemia-induced dopamine release in the corpus striatum of the rat: A study using cerebral microdialysis. Anesthesiology 79:827-835.

Kowalski J, Kwan HH, Prionas SD, Allison AC, Fajardo LF. 1992. Characterization and applications of the disc angiogenesis system. Exp Mol Pathol 56:1-19.

Kramer MS, Vinall PE, Katolik LI, Simeone FA. 1996, Comparison of cerebral blood flow measured by laser-Doppler flowmetry and hydrogen clearance in cats after cerebral insult and hypervolemic hemodilution. Neurosurgery 38:355-361.

Kubota T, Mahler CM, McTiernan CF, Wu CC, Feldman MD, Feldman AM. 1997. End-systolic pressure-dimension relationship of in situ mouse left ventricle. J Mol Cell Cardiol l30:357-363.

Kuluz JW, Prado R, Chang J, Ginsberg MD, Schleien CL, Busto R. 1993. Selective brain cooling increases cortical cerebral blood flow in rats. Am J Physiol 265(Pt 2):H824-H827.

Kushner RF, Schoeller DA. 1986. Estimation of total body water by bioelectrical impedance analysis. Am J Clin Nutr 44:417-424.

Lichtenberg J, Hansen CA, Skak-Nielsen T, Bay C, Mortensen JT, Binderup L. 1997. The rat subcutaneous air sac model: A new and simple method for in vivo screening of antiangiogenesis. Pharmacol Toxicol 81:280-284.

Lindauer U, Villringer A, Dirnagl U. 1993. Characterization of CBF response to somatosensory stimulation: Model and influence of anesthetics. Am J Physiol 264(Pt 2):H1223-H1228.

Lloyd PG, Yang HT, Terjung RL. 2001. Arteriogenesis and angiogenesis in rat ischemic hind limb: Role of nitric oxide. Am J Physiol Heart Circ Physiol 281:H2528-H2538.

Lukaski HC, Bolonchuk WW, Hall CB, Siders WA. 1986. Validation of tetrapolar bioelectrical impedance method to assess human body composition. J Appl Physiol 60:1327-1332.

Maeda K, Hata R, Hossmann KA. 1999. Regional metabolic disturbances and cerebrovascular anatomy after permanent middle cerebral artery occlusion in C57Black/6 and SV129 mice. Neurobiol Dis 6:101-108.

Mathers P, Goode T, Pisacreta D, Motzel S. 2000. Evaluation of multifrequency bioelectrical impedance analysis in non-human primates. (Abstract). Contemp Top 39:61.

Miyamoto, MI, Kim CS, Guerrero JL, Rosenzweig A, Gwathmmey JK, Hajjar RJ. 1999. Ventricular pressure and dimension measurements in mice. Lab Anim Sci 49:305-307.

Moghaddam B, Bunney BS. 1989. Ionic composition of microdialysis perfusing solution alters the pharmacological responsiveness and basal outflow of striatal dopamine. Neurochemistry 53:652-654.

Moses MA, Langer R. Inhibitors of angiogenesis. Biotechnology 9:630-634.

Nawashiro H, Brenner M, Fukui S, Shima K, Hallenbeck JM. 2000. High susceptibility to cerebral ischemia in GFAP-null mice. J Cerebr Blood Flow Metab 20:1040-1044.

Okamota H, Meng W, Ma J, Ayata C, Roman RJ, Bosnjak ZJ, Kampine JP, Huang PL, Moskowitz MA, Hudetz AG. 1997. Isoflurane-induced cerebral hyperemia in neuronal nitric oxide synthase gene deficient mice. Anesthesiology 86:875-884.

Olde Rikkert M, Deurenber P, Jansen R, Vahof MA, Hoefnagels W. 1997. Validation of multi-frequency bioelectrical impedance analysis in detecting changes in fluid balance of geriatric patients, J Am Ger Soc 45:1345-1351.

Orlandi C, Paskins-Hurlburt AJ, Hollenberg NK. 1986. The microvascular response to growth factors in the hamster cheek pouch. Basic Res Cardiol 81:238-243.

Passaniti A, Taylor RM, Pili R, Guo Y, Long PV, Haney JA, Pauly RR, Grant DS, Martin GR. 1992. A simple quantitative method for assessing angiogenesis and antiangiogenic agents using reconstituted basement membrane, heparin, and fibroblast growth factor. Lab Invest 67:519-528.

Sato A, Uchida S, Yamauchi Y. 1994. A new method for continuous measurement of regional cerebral blood flow using laser Doppler flowmetry in a conscious rat. Neurosci Lett 175:149-152.

Scheltinga MR, Helton WS, Rounds J, Jacobs DO, Wilmore DW. 1991. Impedance electrodes positioned on proximal portions of limbs quantify fluid compartments in dogs. J Appl Physiol 70:2039-2044.

Schmidt U, del Monte F, Miyamota MI, Matsui T, Gwathmey JK, Rosenzweig A, Hajjar FJ. 2000. Restoration of diastolic function in senescent rat hearts through adenoviral gene transfer of sarcomplasmic reticulum Ca2+-ATPase. Circulation 101:790-796.

Short CE. 1987. Inhalant Anesthetics in Principles and Practices of Veterinary Anesthesia. Baltimore: Williams & Wilkins. p 72-73, 84-86.

Skarphedinsson JO, Harding H, Thoren P. 1998. Repeated measurements of cerebral blood low in rats: Comparisons between the hydrogen clearance method and laser Doppler flowmetry. Acta Physiol Scand 134:133-142.

Skjeldal S, Nordsletten L, Kirkeby OJ, Grogaard B, Bjerkreim I, Mowinckel P, Torvik A, Reikeras O. 1993. Perfusion in the anterior tibial muscle measured by laser Doppler flowmetry after graded periods of hind limb ischemia in rats. Int J Microcirc Clin Exp 12:107-118.

Stanton CA, Hama DW, Johnson DE, Fettman JM. 1992. Bioelectrical impedance and zoometry for body composition analysis in domestic cats. Am J Vet Res 53:251-157.

Suzuki H, Wilmore DW. 1993. Body composition by bioelectrical impedance analysis during surgical injuries. (Abstract). J Jpn Surg Soc 94(Suppl):P214.

Takeshita S, Isshiki T, Ochiai M, Eto K, Mori H, Tanaka E, Umetani K, Sato T. 1998. Endothelium-dependent relaxation of collateral microvessels after intramuscular gene transfer of vascular endothelial growth factor in a rat model of hindlimb ischemia. Circulation 98:1261-1263.

Taniyama Y, Morishita R, Aoki M, Nakagami H, Yamamoto K, Yamazaki K, Matsumotao K, Nakamura T, Kaneda Y, Ogihara T. 2001. Therapeutic angiogenesis induced by human hepatocyte growth factor gene in rat and rabbit hind limb ischemia models: Preclinical study from treatment of peripheral arterial disease. Gene Ther 8:181-189.

Taniyama Y, Morishita R, , Hraoka K, Aoki M, Nakagami H, Yamazaki K, Matsumotao, K, Nakamura T, Kaneda Y, Ogihara T. 2001. Therapeutic angiogenesis induced by human hepatocyte growth factor gene in rat diabetic hind limb ischemia model: Molecular mechanisms of delayed angiogenesis in diabetes. Circulation 104:2344-2350.

Verbeeck RK. 2000. Blood microdialysis in pharmacokinetic and drug metabolism studies. Adv Drug Del Rev 45:217-228.