Department of Molecular and Cellular Physiology, University
of Cincinnati, Cincinnati, Ohio 45267
The
development and widespread use of genetically altered mice to study the
role of various proteins in biological control systems have led to a
renewed interest in methodologies and approaches for evaluating
physiological phenotypes. As a result, cross-disciplinary approaches
have become essential for fully realizing the potential of these new
and powerful animal models. The combination of classical physiological
approaches and modern innovative technology has given rise to an
impressive arsenal for evaluating the functional results of genetic
manipulation in the mouse. This review attempts to summarize some of
the techniques currently being used for measuring cardiovascular,
renal, and pulmonary variables in the intact mouse, with specific
attention to practical considerations useful for their successful implementation.
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INTRODUCTION |
THE ADVENT OF MOLECULAR TECHNIQUES to
produce targeted gene mutations in the mouse has opened nearly
unlimited opportunities for studying the physiological and
pathophysiological role of almost any functional or regulatory protein
in the intact animal. Mouse models with genetic manipulations
specifically designed to investigate specific proteins are being
developed at a remarkable rate, and the availability of these
"designer mice" has opened avenues of investigation that would have
once been considered implausible. Along with these genetic advances,
considerable progress has been made in our ability to evaluate the
resulting phenotypes at the tissue, organ, and whole animal level.
These advances are evidenced by an ever increasing number and
sophistication of studies that evaluate physiological behavior in
intact mice, which have been made possible by a combination of new
computer and electronic technologies, investigator ingenuity, and
perseverance and not a small amount of mental and manual dexterity.
Commensurate with this progress, there have been a plethora of
illuminating review articles summarizing techniques and methodologies
that have been developed for evaluating function across a wide range of
physiological disciplines, including cardiovascular, renal, pulmonary,
behavioral, neuro-, and electrophysiology (14, 17, 29, 45, 46,
56, 63, 69, 70, 91). It is not the goal of this review,
therefore, to provide yet another synopsis of currently available
approaches, but rather to provide a practical guide to the use and
implementation of these methodologies, primarily for those
investigators who may not have extensive experience with in vivo
experimentation in the mouse. The ubiquitous nature of new transgenic
approaches has, after all, resulted in an inevitable blurring of the
once bold line between molecular biology and classical physiology. By
necessity, investigators wishing to take advantage of these powerful
tools must become at least minimally versed in both disciplines. Because the practical experiences in our own laboratory are primarily centered on cardiovascular, renal, and pulmonary studies, this review
shall focus on these areas of investigation and will necessarily reflect our understanding of the implementation of these techniques. It
is relevant to point out, as many others have, that the mouse is not
simply a small rat, and special care must be taken to resist the urge
to directly compare data from mice to that from rats. Oddly enough, in
our experience, the mouse seems to more closely resemble a small rabbit
in terms of obvious functional characteristics; compared with the rat,
it is somewhat fragile, difficult to maintain blood pressure under
anesthesia, and has a labile blood pressure that is difficult to
elevate experimentally by either acute or chronic intervention.
There are essentially two categories of investigation that are most
commonly used in the phenotypic analysis of newly generated mouse
models: chronic, often noninvasive (or minimally invasive) screening
for functional variations, and acute instrumentation for in-depth
evaluation of specific physiological variables. Examples of chronic
approaches include echocardiography, tail-cuff blood pressure
evaluation, renal balance studies, and whole animal plethysmographic evaluation of pulmonary function. In addition, there has been important
recent progress in the area of chronic instrumentation and the use of
telemetric techniques for evaluating function in a relatively
undisturbed animal. Acute approaches can be categorized into in vivo
procedures, such as cardiac catheterization, regional blood flow
measurements, renal clearance and micropuncture studies, and airway
pressure-flow measurements, as well as ex vivo procedures, such as
Langendorff and isolated working heart preparations (37), isolated smooth muscle preparations (79), and isolated
lung preparations (67). Because the requirements for
adequate phenotypic evaluation of mutant mouse strains is becoming more
rigorous in the eyes of many reviewers, this article will describe the
various screening methods often used to initially evaluate organ
function in mice, but will focus more intently on the more analytic
techniques used to confirm various phenotypic traits.
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ANESTHESIA |
The choice of anesthesia for use in mice is a crucial decision and
depends largely on the type of experiment being performed as well as
the personal preference and experience of each investigator. Variables
to be considered are background strain of mouse, duration of the
procedure, whether the procedure is terminal or recovery in nature, and
the physical constraints of the planned instrumentation. Anesthetic
regimens are of two types, injectable and inhaled, and the various
compounds used in mice have been extensively reviewed (69). The most commonly used anesthetics in mice include
the injectable agents avertin, pentobarbital, inactin
(thiobutabarbital), chloralose-urethane, ketamine (usually combined
with other agents such as acepromazine, xylazine, and/or diazepam), and
inhaled agents isoflurane, methoxyflurane, and halothane. We have
adopted the use of several anesthetics in our laboratory and the choice depends on several factors. For long, nonrecovery procedures, such as
cardiac catheterization or renal micropuncture, we typically use a
combination of ketamine and inactin given as separate intraperitoneal injections: 2 µl/g body wt of 25 mg/ml ketamine given first, followed by 2 µl/g body wt of 50 mg/ml inactin. This combination allows for
quick induction and fairly prolonged action with minimal
supplementation required. The cardiodepressor effects are mild as
evidenced by mean arterial pressure in the 80-90 mmHg range and
left ventricular dP/dt ~9,000 mmHg/s under basal
conditions (58, 59); these values are only slightly lower
than those observed in awake mice (66). The advantage of
using both agents in a long procedure is that the ketamine-inactin
combination will initially produce a deeper anesthesia that permits
relatively invasive procedures. Then as the shorter acting ketamine
begins to wear off, the level of anesthesia can be carefully titrated
via intravenous supplementation and during constant monitoring of heart
rate and blood pressure. It should be recognized that inactin does not
have the very long duration of action that is seen in the rat and
therefore must be supplemented regularly; nonetheless, it is generally
accepted that its use for recovery surgeries should be avoided. For
nonrecovery procedures, a tracheotomy is usually performed using a
short length of PE-90 tubing to ensure airway patency, but the animal
is usually allowed to breath spontaneously without the aid of a
ventilator. However, it is fairly common to provide anesthetized mice
with a stream of 100% O2 to prevent hypoxia. It must also
be noted that anesthetized mice are particularly vulnerable to
hypothermia and body temperature must be constantly monitored and
carefully regulated. We monitor and maintain temperature via a rectal
thermistor probe and a feedback-controlled, warmed surgical table
(Vestavia Scientific, Birmingham, AL). Supplemental heat is often
provided using a heat lamp to ensure even warming.
We recently began to explore the use of inhaled anesthetics (isoflurane
in our case) for prolonged analytic procedures. The advantages of
isoflurane are that it preserves sympathetic vasomotor activity, has
minimal cardiodepressor effects, and allows very careful,
minute-to-minute control of the anesthetic plane. By way of comparison,
left ventricular dP/dt measured under isoflurane averages
~13,000 mmHg/s and mean arterial pressure is usually ~90-100
mmHg (unpublished observations). The disadvantages of isoflurane are
that it requires expensive equipment, can be physically cumbersome,
especially when the procedure requires extensive
instrumentation and multiple changes in animal posture, and the
depth of anesthesia can be volatile if its administration is not finely
and carefully controlled, often through the use of mechanical
ventilation. We attempted to combine isoflurane with other agents, such
as ketamine or buprenorphine, to help stabilize the anesthetic plane,
but many of the benefits of inhaled anesthesia are diminished by this approach. In our experience, isoflurane anesthesia can be quickly induced by manually restraining the animal and placing its head in a
mask fashioned from a 50-ml Falcon tube through which a steady stream
of 4% isoflurane is introduced. After induction of deep anesthesia,
usually within 30 s, the animal is removed from the mask and
quickly orally intubated, and isoflurane is provided at 2.25% via
spontaneous breathing or through a ventilator. Oral intubation using a
20-gauge stainless steel or Teflon needle can be accomplished by
illuminating the ventral surface of the neck with a bright light source
and then visualizing the vocal cords (4) or, more simply,
by quickly exposing the trachea via a small neck incision and advancing
the cannula under direct visualization of the airway. With minimal
practice, intubation can be accomplished in less than 30 s and a
steady flow of anesthetic can be reestablished well before the animal
begins to awaken. Of course, inhaled anesthetics are also ideal for
recovery procedures because they are so well tolerated and have fast
recovery times. After cessation of isoflurane, mice usually begin to
regain consciousness within 2 min and are fully ambulatory in 5-10
min. These properties make it an ideal choice for the implantation of
indwelling catheters or telemetry probes or for surgical manipulations,
such as transthoracic aortic banding or coronary artery ligation.
Furthermore, because of its minimal cardiodepressant effects and quick
induction and quick recovery time, the use of isoflurane is a preferred
choice for brief analytic procedures that require sedation, such as
echocardiography (80).
We also used ketamine mixtures in our lab for both recovery procedures
and nonrecovery analytic procedures. For mice, a mixture of 67 mg/ml
ketamine, 3.3 mg/ml xylazine, and 1.7 mg/ml acepromazine given
intraperitoneally at a dose of 1.5 µl/g body wt seems to work quite
well. This cocktail is made by mixing standard preparations as follows:
8 ml of 100 mg/ml ketamine, 2 ml of 20 mg/ml xylazine, and 2 ml of 10 mg/ml acepromazine. This mixture has moderate cardiodepressant effects,
but is well tolerated and produces a surgical plane of anesthesia for
~1 h when given as a single intraperitoneal injection. We commonly
use this preparation for recovery surgeries such as catheter
implantation and renal artery clipping. It provides ample time to
complete the surgery, permits free manipulation of the animal's
posture and position, and the animal generally is fully ambulatory
within 2 h. In addition, we used this anesthetic for the analysis
of airway reactivity in nonrecovery experiments. Unlike the
barbiturates, the ketamine mixtures appear to preserve more of the
airway responsiveness to bronchoconstrictor challenges such as
acetylcholine administration.
For very short procedures, we often used avertin, which has the
advantage of having very fast induction and recovery times. Induction
of a surgical plane of anesthesia usually occurs within 2-3 min of
injection and can last 15-30 min; animals also awake quite
suddenly, becoming ambulatory within minutes of the initial signs of
recovery. The preparation, which is no longer available commercially,
is made as a 100% stock solution by mixing 10 g of
tribromoethanol with 10 ml of tertiary amyl alcohol. This stock is then
diluted to 2.5% and given as an intraperitoneal injection at a dose of
15-17 µl/g body wt (44). Both solutions should be
stored in the dark at 4°C, and, in our experience, the 100% stock is
stable for at least 6 mo, but the 2.5% stock should be prepared fresh
at least every 2 wk. We often used smaller doses (10-12 µl/g
body wt) as a sedative for performing echocardiographic measurements,
but even at these lower doses, avertin is cardiodepressant, often
yielding heart rates of <400 beats/min and blood pressures of
~70-80 mmHg. Avertin, which is most often used for embryo
transfer, can be an effective anesthetic for brief recovery surgical
procedures, but, because of its short duration of action and rapid
recovery period, it should be used with caution. Supplements can be
given, but there are reports of acute necrotic and inflammatory changes with higher doses and increased mortality with repeated administrations (95). In any case, investigators should keep in mind that
the choice of anesthesia should depend not only on the procedure being performed, but also on the organ system or systems being studied. For
instance, urethane is often used in the rat by neuroscientists because
it permits maintained neural activity and blood pressure, but it can be
a poor choice for renal investigations because of intraperitoneal
toxicity and derangements in renal hemodynamics and fluid handling
(38, 75).
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GENERAL PROCEDURES |
Instrumentation.
It has been fortuitous, but not altogether coincidental, that the
advent of genetically altered mice as models for physiological investigation has corresponded with significant advancements in electronics, instrumentation, and computer technology. Microchip-based instrumentation has permitted the miniaturization of sensors and transducers to a size and performance level suitable for the mouse. In
addition, computer technology and digital recording equipment, along
with significant advances in data archival media, have allowed for
greater accuracy, ease, and dependability in recording equipment.
In general, there have been two obstacles to overcome in adapting
existing methodologies for use in the mouse: the size, which is about
one-tenth the size of the traditional rat model, and the speed or
frequency-response requirements, which are up to twofold greater than
in the rat. Equipment manufacturers, having been convinced of the
importance and endurance of mouse models, have for the most part been
responsive to the particular challenges presented by an animal that
weighs only 30 g. For example, since first being used in the
intact mouse, high-fidelity microtip transducers from Millar
Instruments (Houston, TX) have been reduced in size by 20% (from 2 Fr.
to 1.4 Fr.), making left ventricular pressure measurements routine
(59). Transit-time flow probes (Transonics Systems) have
been miniaturized to a point where they are capable of accurately
measuring flow in vessels as small as 0.25 mm in diameter, such as the
renal artery (33), and further miniaturization is on the
horizon. Telemetric implants have even been miniaturized to the point
where heart rate, temperature, and blood pressure can be monitored
continually in unrestrained mice for months at a time (5,
24). Along with advancements in sensors, there have been radical
improvements in data recording and analysis equipment, and there are a
number of hardware/software packages on the market that greatly
simplify and improve the recording of physiological signals. In our
laboratory, we primarily use a PowerLab System (ADInstruments, Grand
Junction, CO), which permits recording at very high sampling speeds and
has a wide array of signal recording, conditioning, and analysis
options. It is important to note that the sampling speed and the
frequency-response characteristics of an entire recording system must
be considered carefully. As an example, consider the measurement of
left ventricular pressure (see below for more detail). Under maximally
stimulated conditions, left ventricular dP/dt can be
as high as 30,000 mmHg/s at a heart rate of ~700 beats/min. Signals
that occur this quickly cannot be monitored with conventional
fluid-filled catheters, because the signal is dampened by the
necessarily small lumen diameter. In addition, conventional paper
strip-chart recorders are unable to accurately follow signals at this
high rate. To achieve sufficient frequency-response characteristics,
left ventricular pressure measurements are therefore typically made
using Millar Mikro-Tip pressure transducers coupled to a computer-based
recording system operating at a sampling speed of 1,000 or even 2,000 Hz (59). A sample tracing of left ventricular pressure and
dP/dt is shown in Fig. 1.
Figure 1, top, shows the actual data points that describe the ventricular pressure pulse and illustrates the importance of a high
frequency-response, both in measuring and recording equipment. At
baseline function (Fig. 1A, dP/dt 
10,000
mmHg/s), the majority of the pressure increase during systole occurs in ~10 ms and it is clear that a sampling rate of 1,000 Hz is adequate. At extremely elevated performance levels (Fig. 1B,
dP/dt 30,000 mmHg/s), the majority of the pressure
increase occurs in <5 ms, and a higher sampling rate is necessary to
faithfully record this waveform. By contrast, dP/dt values
in the rat range from ~7,000 to perhaps 16,000 mmHg/s. Although
current technology allows for fluid-filled catheter systems that are
nicely responsive (frequency-response curves that are flat to perhaps
150 Hz), these figures also demonstrate that only high-fidelity
microtip transducers, with a flat frequency response up to 10,000 Hz
(i.e., Millar), are sufficient for evaluating left ventricular pressure
in mice. For other pressure measurements, we found fluid-filled systems
to be more than adequate.
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Fig. 1.
Sample tracing of left ventricular (LV) pressure (top)
and dP/dt (bottom) during baseline conditions
(A) and infusion of dobutamine at a dose of 32 ng · g body wt 1 · min1
(B). Pressure tracings are displayed as discrete points to
illustrate the importance of sampling rate: at baseline the systolic
pressure rise is largely complete within 13 ms, but during dobutamine
treatment, the pressure rise is complete in ~5 ms. This is reflected
in peak dP/dt values in the bottom tracings.
Pressure measurements were made using a Millar 1.4-Fr. Mikro-Tip
transducer advanced to the LV through the right carotid artery; data
were recorded using an ADInstruments PowerLab 4SP.
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Vessel isolation and cannulation.
Many, if not most, of the techniques used to evaluate cardiovascular,
renal or pulmonary function involve the surgical isolation and
cannulation of an artery and or vein and the techniques used do not
vary substantially from those used in larger animals. The right tools
are essential for these procedures, and great care must be taken to
prevent even slight blood loss: the total blood volume of a mouse is
2-3 ml and the loss of even 100 µl of blood can represent a
significant hemorrhage. We found use of a low-power binocular
dissecting microscope to be essential for procedures, and adding ×0.5
objectives can increase the field of view as well as the working
distance. Dissecting tools include several pairs of very fine forceps
(i.e., Dumont #5, straight or angled), small dissecting scissors, and a
set of Vannas scissors. For fluid-filled catheters, we found it
convenient to pull very fine cannulas over an open flame from
large-bore, thick-walled polyethylene tubing. With the use of this
process, the tubing is placed just above the hot flame until it becomes
molten and then it is removed and quickly stretched to produce a fine
capillary. With a little practice, this approach can be used to make
catheters of almost any dimension. To minimize the dead space for
venous catheters, we start with polyethylene tubing with an outer
diameter (OD) of 1/4 in. and a wall thickness (WT) of
in. (6.4 mm OD × 1.6 mm WT, Fisher Scientific).
Conversely, to increase the relative lumen size for arterial catheters,
we start with polyethylene tubing with an OD of 
or
1/2 in. (9.5 or 12.7 mm) and a WT of in. The
resulting catheter has a relatively large ID-to-OD ratio and therefore
better frequency-response characteristics than conventional PE tubing
(i.e., stretched PE-10) when measuring arterial blood pressure. With
this approach, we have been able to fashion catheters as small as 100 µm OD and have managed to insert up to four catheters into a single
femoral vein.
For actual cannulation, the vessel is carefully isolated from
surrounding tissue and associated vessels using two pairs of fine
forceps. We found it useful to polish the tips of Dumont #5 forceps to
remove burrs and to slightly blunt them to prevent accidental puncture
of the vessel (the veins are especially delicate and can be easily
torn). Once isolated, three loops of 7-0 silk suture are placed
around the vessel; the distal-most ligature is tied to prevent back
flow of blood and enough tension is placed on the proximal ligature so
as to temporarily occlude flow. The center ligature is loosely knotted
and is to be used to secure the catheter in place and prevent leakage
when the proximal tie is released. Once blood flow is occluded, a short
longitudinal incision is made in the vessel and the cannula is
introduced. Alternatively, some investigators use a 23- or 25-gauge
hypodermic needle, bent at an angle at the tip, to puncture the vessel
and to use as an introducer. Once the cannula is in place, the central ligature is tightened and the tension is removed from the proximal tie;
the catheter can then be advanced further and secured into place. All
surgical incisions are closed using cyanoacrylate adhesive, even in
nonrecovery procedures, to prevent evaporative fluid loss and tissue desiccation.
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CARDIOVASCULAR MEASUREMENTS |
It is often useful to monitor various cardiovascular parameters
over long periods of time, weeks to months, without surgical interventions. We have used three well-described techniques to accomplish this: tail-cuff measurement of blood pressure,
echocardiography, and exercise tolerance. In addition, radiotelemetry
devices have recently been developed that are small enough to permit
measurement of a variety of variables such as electrocardiogram,
temperature, activity, and blood pressure. Finally, for in-depth
analysis of cardiovascular function in vivo, high-fidelity measurements
of left ventricular function, cardiac output, and regional blood flow
can be made in acute and, in some cases, chronic preparations.
Tail-cuff pressure.
The measurement of systolic pressure through tail sphygmomanometry has
been a standard technique for the long-term evaluation of blood
pressure in the rat and has been applied to the mouse under a variety
of experimental paradigms (7, 42, 85). Although similar
measurements in the mouse are technically more challenging, updated
instrumentation, using computer-controlled pulse detection and
data-acquisition technology, was recently developed and validated and
provides a practical approach for these measurements in the mouse
(53). We are currently using an integrated system
(Visitech, Apex, NC) that permits measurement of tail-cuff pressure on
four mice simultaneously and requires a minimal training period
(4-7 days). Using this system, we can perform measurements on
12-16 mice per hour. As with any tail-cuff approach, measurements
must be obtained over an extended period of days (usually ~5-7
days, in addition to the training period), to obtain a reasonable
determination of blood pressure in a group of animals. As an
illustration of the effectiveness of the tail-cuff approach in the
mouse, we examined the development of hypertension in mice that were
fed NG-nitro-L-arginine methyl ester
(to inhibit nitric oxide production) over a 30-day period and the
results are shown in Fig. 2. We have used
this system extensively and have found that it can generally discern
differences in blood pressure as low as 10-15 mmHg.
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Fig. 2.
Systolic blood pressure measured using Visitech Systems
blood pressure monitor. Ten measurements were taken each day, and the
average measurement for each mouse was reported. Mice were trained in
the restraining device for 5 days before the initiation of the study,
and each episode of 10 measurements was preceded by 10 preliminary
measurement, which were discarded. One group of animals
(n = 6) was treated with 0.25 and 0.5 mg/ml
NG-nitro-L-arginine methyl ester
(L-NAME) in their drinking water, as indicated, whereas the
other received distilled water.
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Although the tail-cuff method provides a simple estimate of arterial
blood pressure, it has a number of disadvantages that can limit its
usefulness. Primary among these is the stress it produces in the
subject animals. Although acclimating the animals to the instrument
helps to some degree, it must be recognized that these blood pressure
measurements are taken under some degree of anxiety and stress. This is
most easily evidenced by the high heart rates measured by tail cuff: as
high as 650-700 beats/min, compared with ~500 beats/min in the
resting animal. Unlike the rat, which usually becomes quite comfortable
in restraint after moderate training, the stress level in mice (as
reflected in heart rate) remains elevated even after several weeks of
training. Fortunately, the available evidence indicates that the blood
pressure measurements are less affected by the imposed stress
(61). Measurements of tail-cuff pressure are also
notoriously variable from reading to reading, and this problem can be
exacerbated by the software detection algorithms designed to
automatically find the systolic pressure end point during cuff
inflation. To explore this potential problem we recorded tail-cuff
measurements while simultaneously monitoring intra-arterial blood
pressure using implanted telemetry devices; a sample tracing is shown
in Fig. 3. When the systolic end point is
clearly discernible and is without movement artifact, there is very
good agreement between arterial pressure and tail-cuff pressure and
most of the variation is due to actual moment-to-moment variation in
the subject's blood pressure. However, investigators should be
cautioned that although the automatic detection algorithms are
generally quite good when the animal is quiet, movement artifacts, which occur quite often even in trained animals, can result in erroneous measurements that should be discarded. It is therefore important to visually inspect the pulse tracings while making these
measurements and to make note of those measurements that do not conform
to preestablished criteria. It is also important to take multiple
readings to compensate for the overall variability (see Fig. 2 legend).
Finally, it is interesting to note that the tail-cuff pressure
generally correlates well with mean, rather than systolic, pressure, a
finding that is consistent with previous reports (53). We
presume this observation to be due to a significant pressure gradient
between the carotid artery measurement site and the tail. This would
not be surprising given the relatively small caliber (~0.5-1.0
mm) of the aorta. Thus, as in the rat, the tail-cuff method can be a
valuable approach for screening and monitoring changes in blood
pressure over extended periods of time if used with proper caution and
diligence. However, as is also the case in the rat, the limitations of
the tail-cuff approach prescribe that independent verification of blood
pressure phenotype by an alternative method be obtained.
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Fig. 3.
Tracing of tail-cuff pressure (A; bold line)
and tail pulse (B) from a mouse using the Visitech blood
pressure monitor, superimposed over a simultaneous recording of
intra-arterial blood pressure obtained from a Data Sciences PA-C20
blood pressure telemetric implant (A; fine line). Note that
the end point of the tail cuff measurement, coinciding with the
disappearance of the tail pulse and indicated by the vertical dashed
line, corresponds approximately with mean arterial pressure obtained
from the pressure implant.
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Echocardiography.
In similar fashion to evaluation of tail-cuff pressure over
extended periods of time, it is often useful to evaluate cardiac function over a period of weeks or months, and transthoracic
echocardiography of the mouse heart has become well established as a
method for providing useful indexes of myocardial performance. Use of
M-mode and pulsed-Doppler echocardiography to assess ventricular
function in the mouse has gained widespread use as a noninvasive method for evaluating cardiac function and has been previously reviewed in
detail (12, 45, 46). Although echocardiographic methods in
mice do not differ conceptually from those in rat, actual images are
more difficult to obtain due to the small size, increased rate of
shortening, and increased relative distance of the heart from the chest
wall. As newer, more advanced imaging technology has become available,
spatial and temporal resolution for imaging the mouse heart has become
more and more accurate. Two-dimensionally directed M-mode
echocardiography permits a number of parameters to be determined,
including left ventricular end-diastolic and end-systolic dimension and
posterior and anterior (septal) wall thickness. From these dimensions,
one can calculate variables such as left ventricular fractional
shortening and left ventricular mass. Furthermore, pulsed Doppler
measurements of transvalvular aortic and mitral flow can be used to
obtain ejection phase indexes of systolic and diastolic function. When
combined with heart rate-matched M-mode images, one can calculate a
variety of performance indexes such as aortic acceleration and ejection
time, velocity of circumferential shortening, peak aortic flow
velocity, and early and late diastolic transmitral velocity. Although
Doppler measurements of flow velocity are limited by the ability to
direct the ultrasound beam parallel to flow in the mouse, measurements
such as ejection time and acceleration time can be reliably determined.
Newer generation high-frequency transducers and scanners provide real
time analysis of two-dimensional images with high temporal resolution,
allowing more accurate serial assessment of left ventricular mass
(20).
In our laboratory, mice are sedated with an intraperitoneal injection
of 2.5% avertin (10 µl/g body wt) and the chest is shaved. The
animal is placed in a supine position and warmed using an isothermal
heating pad. Ultrasound studies are performed using an Interspec Apogee
X-200 ultrasonograph and a dynamically focused 9-MHz annular array
transducer with an axial resolution of 0.2 mm. Acoustic coupling is
achieved through a gel-filled offset applied to the shaved chest.
Two-dimensional targeted M-mode studies are taken from either the long
or short axis at the level of greatest left ventricular diameter.
Systolic aortic outflow and diastolic transmitral inflow velocities are
determined from angulated parasternal long-axis views using a pulsed
wave Doppler transducer. Left ventricular function can be determined
under baseline conditions and during 
-adrenergic stimulation by
intraperitoneal injection of isoproterenol. Tracings are recorded on
S-VHS tape, and frozen images are later digitized. Measurements are
made from the digital images using image analysis software (NIH Image),
and three beats are averaged for each measurement. Sample M-mode and
pulsed-Doppler wave forms are shown in Fig.
4. In the M-mode image in Fig.
4A, the anterior and posterior left ventricular walls are
clearly defined and permit reasonably accurate determination of
end-systolic and end-diastolic dimensions and wall thickness. As
illustrated in Fig. 4B, Doppler flow recordings allow
time-based measurements such as heart rate (from R-R interval),
acceleration time, and ejection time, as well as measurement of
flow-dependent variables such as peak aortic velocity.
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Fig. 4.
A: representative M-mode tracing from a
wild-type mouse. AW, anterior LV wall; PW, posterior LV wall; EDD:
end-diastolic dimension; ESD, end-systolic dimension; AWth, anterior
wall thickness; PWth, posterior wall thickness. B: aortic
Doppler waveforms from a wild-type mouse. PaoVel, peak aortic velocity;
AT, acceleration time; ET, ejection time; R-R, R-R interval. Images
were obtained using an Interspec Apogee X-200 utrasonograph and a
dynamically focused 9-MHz annular array transducer with an axial
resolution of 0.2 mm.
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A pressing issue in echocardiographic analysis is the method of
sedation, if any. Numerous studies reported on the effects of various
anesthetics on echo-derived heart function, and several labs reported
studies performed without sedation. A recent study compared
xylazine-ketamine vs. avertin anesthesia on cardiac function in mice
using both echocardiography and closed-chest catheterization (40). The authors reported that compared with avertin,
xylazine-ketamine anesthesia produced severe bradycardia with secondary
effects on loading conditions, and their findings illustrated the need for comprehensive assessment of left ventricular function in the mouse
with specific attention demanded regarding the mode of sedation. Another study compared the effects of halothane vs. ketamine-xylazine (10), which also concluded that echo-derived parameters
are highly sensitive to anesthetic regimen and that halothane is
superior to ketamine-xylazine for obtaining relevant performance
parameters. The feasibility and effectiveness of performing
echocardiography studies in restrained nonanesthetized mice have also
been evaluated (94). In this study, parameters obtained
from awake mice were compared with those from pentobarbital- and
ketamine-xylazine-anesthetized mice. Predictably, findings showed that
both anesthetics result in a marked decrement in heart rate and
performance variables compared with awake mice; it should be noted,
however, that the echo-derived heart rate in awake mice was
substantially higher than heart rates determined in undisturbed animals
by telemetry. In any case, it is clear that the interpretation of
echocardiographic data must be interpreted with caution, including
serious consideration of the anesthetic state of the animal. As with
tail-cuff measurement of blood pressure, it is recommended that
differences in cardiac performance observed with echocardiography be
verified using an alternative methodology (see Cardiac
catheterization).
Exercise tolerance.
Experimental mouse models with deficits in cardiovascular function can
be expected to exhibit an impaired ability to tolerate exercise. It has
been our experience, in fact, that many animal models that appear
phenotypically normal under resting conditions can exhibit deficits in
their ability to tolerate even moderate exercise, and such measurements
can provide a useful integrated index of overall cardiovascular
integrity. To objectively evaluate exercise tolerance in mice, we
combined an exercise treadmill (Omnipacer, Accuscan Instruments,
Columbus, OH) with a custom-designed, personal computer-based detection
system, permitting automatic quantification of an animal's performance
(27). A motor-driven treadmill, with adjustable belt speed
and shock grid, was modified by installing an infrared detection system
consisting of two infrared detectors and logic circuitry assembled on a
printed circuit board. When a mouse running on the treadmill blocks the
infrared sensor, this "failure" produces a signal to the digital
input/output board inside a personal computer, which is then
recorded and displayed in real time by custom software. This program
checks and records the status of the infrared switches (on/off) once
every second to determine the exercise status of the subject. A similar
system was described in detail in a report that evaluated exercise
performance, in conjunction with telemetric recording of heart rate, in
mice overexpressing ventricular myosin regulatory light chain
(24). In our experience, it is important to acclimate the
animals to the treadmill before evaluating performance. However, as the
goal is usually to evaluate basal exercise performance rather than the
response to exercise conditioning, we prefer to keep the acclimation regimen quite mild, usually 15 min at a relatively slow speed (10 m/min) for 2 or 3 days. Our measurement protocol, then, usually consists of a 40-50 min exercise period in which the treadmill is
set at a 7° incline and the speed is increased by 5 m/min every 10 min. This protocol is repeated three times over 5 days.
Telemetry and indwelling catheters.
It is accepted that the most reliable of cardiovascular measurements
are those obtained using radiotelemetry. Although implants suitable for
measuring electrocardiogram, temperature, and activity in mice have
been available for a number of years (24, 48, 49), only
recently has the miniaturization of these devices permitted blood
pressure measurement in the mouse (65). These devices
(Data Sciences International, St. Paul, MN), which enable long-term
continuous recording of blood pressure without restraint, have become
accepted as the gold standard for accurate evaluation of blood pressure
in the intact, conscious animal. Although these implants provide
significant benefit over other methodologies, their use in the mouse
has presented several potential limitations. First, despite being
significantly miniaturized, the transmitters are still relatively
large: ~1 × 1.5 × 2.3 cm in size (~ 2.3 ml) and ~3.5
g, roughly 10-15% of the normal body weight of a mouse. In
earlier applications, the sensing catheters on these devices were
implanted in the abdominal aorta in a nonoccluding fashion, with
intraperitoneal placement of the transmitter, and their recommended use
was limited to 30 g or larger mice. Furthermore, even in these larger mice, the size of the catheter in relation to the abdominal aorta was such that could often occlude flow to the hindquarters, leading to a discouragingly low success rate. To solve these problems, Carlson and Wyss (6) recently published a report
describing carotid cannulation and subcutaneous placement of the
transmitter on the animal's back, obviating the need for the more
invasive abdominal implantation. Using this protocol, these
investigators reported a success rate of >90% in mice as small as
19 g. This technique was further modified so that the transmitter
was placed along the flank of the animal, greatly simplifying the
implantation procedure (5). The advantages of subcutaneous
implantation are such that it is recommended under certain
circumstances for use in rat as well as in the mouse: it is less
stressful surgically and is characterized by a faster return to
presurgical weight and circadian patterns than abdominal placement.
Nonetheless, it is important to note that telemetry studies have
consistently demonstrated that full recovery from anesthesia and
surgery does not occur for 5-7 days, as indicated by the return of
normal circadian rhythms in activity, blood pressure, and heart rate.
Other disadvantages of radiotelemetry are that it is expensive and does
not allow vascular access for the administration of experimental
agents. As an alternative, several investigators have adapted the use
of indwelling catheters and swivel tethering systems to monitor
intra-arterial blood pressure in mice (47, 50, 61, 62). In
this procedure, catheters are implanted in the femoral vessels or
carotid artery and tunneled subcutaneously to the nape and exteriorized
through a spring, which is secured to the animal's back via a Teflon
button. The spring can then be connected to a swivel device at the top
of the cage to allow free movement of the tethered animal. Although
catheter construction varies widely, we found that polyurethane tubing
(Micro-Renathane, 0.25 mm OD, Braintree Scientific, Braintree, MA)
pulled to a small diameter in hot oil is very effective and usually
remains patent for extended periods. With the use of this approach,
stable blood pressure and heart rate measurements have been recorded
for as long as 5 wk (61). Because this technique allows
for continual vascular access and blood pressure monitoring without
disturbance to the animal, it can be used to evaluate a variety of
cardiovascular parameters, such as the sensitivity of blood pressure to
precisely regulated electrolyte intake, baroreflex sensitivity, and
blood pressure and heart rate variability (47, 50).
Cardiac catheterization.
As indicated in an earlier section of this discussion (see
Instrumentation), in vivo catheterization of the left
ventricle in the closed-chest mouse has become commonplace and is a
well accepted standard for evaluating cardiac performance in
genetically manipulated mice. Due to the small size and
frequency-response requirements of the murine heart, a Millar Mikro-Tip
transducer is the only suitable alternative for measuring left
ventricular pressure in the mouse. Although these catheters are
somewhat costly, they have in our experience proven to be quite rugged,
and with proper care can be used for extended periods of time and for
hundreds of experiments. It is important to note that these transducers are exquisitely sensitive and can be significantly influenced by
ambient conditions such as temperature, composition, and viscosity of
surrounding fluid (i.e., blood vs. saline), and even light. For this
reason, we follow a strict set-up and calibration procedure when
performing left ventricular pressure experiments. First, as emphasized
by the manufacturer, the catheter is presoaked in saline for at least
30 min before implantation; we presoak in a darkened cuvette at 37°C.
Just before implantation, the bridge amplifier is balanced and
calibrated using the built-in electronic calibration feature of the
Millar control unit. The catheter is then introduced into the carotid
artery as described previously and advanced to the heart and across the
aortic valve under the guidance of the online pressure signal (this can
usually be accomplished without difficulty). At the end of the
experiment, the Millar catheter is withdrawn from the carotid artery
and a small pool of blood is allowed to form in the neck cavity of the
mouse; the transducer tip is immersed just below the surface of this
pool to obtain a value at atmospheric pressure in a fluid environment (temperature and viscosity) that most closely resembles that within the
heart chamber; this pressure value is defined as zero. Importantly, this value can differ from the value obtained in saline at the beginning of the experiment by several millimeters of mercury, a
difference that can be crucial when trying to evaluate and interpret variables such as left ventricular end-diastolic pressure. Finally, the
catheter tip is placed in a closed tube of warmed saline that is
connected to a mercury manometer and pressurized to a series of known
pressures. In this manner, a "wet calibration curve" can be
recorded and appended to the experimental tracings. The digital
data-acquisition system (in our case the PowerLab System) permits the
entire pressure and dP/dt recordings obtained during the
experiment to then be recalibrated using the values obtained at the end
of the experiment.
Although measurements of left ventricular pressure have proved to be
extremely valuable in assessing cardiac performance in genetically
altered mice, the analysis of pressure alone cannot account for
potential differences in loading conditions of the heart from one mouse
to the next. To reliably evaluate cardiac contractility, several groups
of investigators sought to simultaneously evaluate left ventricular
pressure and volume to obtain load-independent indexes of contractile
function. For example, we reported the use of echocardiography in
conjunction with simultaneous recording of left ventricular pressure to
derive end-systolic pressure-dimension relationships in intact mice
(13). Although this approach proved effective, it did not
permit continuous beat-to beat evaluation of pressure-volume
relationships and the image-based data analysis was extremely labor
intensive. Investigators also successfully used sonomicrometry to
evaluate cardiac dimension during left ventricular pressure
measurements (18, 54). In this procedure, piezoelectric
crystals are implanted on or within the myocardium in an open-chest
preparation, and measurements of left ventricular dimension are
recorded simultaneously with pressure recordings via a Millar catheter
placed in the left ventricular cavity. As demonstrated in one recent
study (18), this technique can be quite effective when
used with two pairs of crystals and can yield reasonable
pressure-volume relationships. However, this technique has the distinct
disadvantage of being very technically demanding and highly invasive,
requiring significant intrathoracic manipulation as well as open-chest
measurements. This is probably reflected in low values of cardiac
output and ejection fraction (<5 ml/min and 30%, respectively). The
most promising and productive technique for evaluating pressure-volume
relationships in mice has been by conductance measurement using a
Millar 1.4 pressure catheter with four integrated platinum electrodes.
This method, first reported in the mouse by Georgakopoulos et al.
(31), permits the generation of an instantaneous signal
for left ventricular volume that is based on the time varying
conductance measured by the two pairs of electrodes within the left
ventricular chamber. Essentially, changes in the electric field
generated by one pair of electrodes during chamber filling and ejection
is sensed as a change in voltage in the other pair of sensing
electrodes. Signal conditioning and processing is accomplished through
specialized instrumentation, such as the Aria-1 Conductance System from
Millar Instruments. This technique can be used in the closed- or
open-chest animal and has the advantages of being no more technically
demanding than standard left ventricular catheterization. Disadvantages include difficulty in calibrating the volume signal and the need to
account for the component of the conductance signal that is not
dependent on chamber dimension, that is, the parallel conductance due
to surrounding structures (i.e., myocardium). There have been several
reports that explore the use of this technique in the mouse in
considerable depth and the reader is directed to these references for
more technical consideration of the relevant issues (22, 23, 30,
31, 93). Sample pressure-volume traces from an
isoflurane-anesthetized animal using the conductance method are shown
in Fig. 5.
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Fig. 5.
Pressure-volume relationship under baseline conditions
(fine tracing) and during dobutamine infusion (bold line) obtained from
preload reduction by applying a positive end-expiratory pressure with
the ventilator. The resulting increase in alveolar pressure transiently
decreases blood flow through the lung and thereby decreases venous
return to the left heart. Note that -adrenergic stimulation shifts
the end-systolic relationship upward and to the left. Inset:
actual tracings of pressure (top) and volume
(bottom). Data were collected using a Millar 1.4-Fr.
pressure-conductance catheter and Aria-1 conductance system. The volume
signal was calibrated using lucite chambers of known volume filled with
fresh heparinized whole blood, and the parallel conductance was
determined using the hypertonic saline injection method
(30).
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Because cardiac contractility can be significantly influenced by
changes in heart rate, it can often be difficult to compare cardiac
performance in animals that have different resting heart rates.
Furthermore, in our experience, it is not unusual for genetically manipulated animals to exhibit chronotropic as well as inotropic phenotypes. Thus, in many instances it can be useful, if not essential, to electrically pace the heart in an in vivo preparation. To accomplish this, a bipolar pacing wire with a blunted tip is inserted into the
right jugular vein and advanced to the right atrium. This pacing
electrode is then connected to a suitable stimulator, and a normal
sinus rhythm can be produced at any rate above the normal resting heart
rate. In this manner, ventricular performance in animals with differing
resting heart rates can be compared by pacing them at a common
frequency (15). In addition to providing a means to
normalize heart rate between animals, atrial pacing allows for the
determination of valuable force-interval relationships in mouse models
of cardiac dysfunction. The most straightforward of these is the
force-frequency relationship, which has been interpreted in terms of
variable amounts of Ca2+ made available to the contractile
myofilaments as a function of the interval between beats.
Blood flow and cardiac output.
Evaluation of regional blood flow and cardiac output is also an
important component for thorough analysis of any cardiovascular phenotype, and a variety of approaches and techniques has been adopted
for use in the mouse. In a comprehensive series of studies in a mouse
model overexpressing atrial natriuretic factor, direct intraventricular
injection of radioactive microspheres was used to estimate blood
volume, cardiac output, and regional blood flows in intact, awake mice
(1, 2). Although effective, this method has the
disadvantage of yielding measurements at only one time point. In
addition, a number of attempts has been made to use indicator dilution
methods for analyzing cardiac output in mice, but this methodology is
difficult to apply in small animals for a variety of reasons, including
loss of diffusible indicator in the pulmonary vascular circuit, lack of
vascular access to the pulmonary artery, and limited capacity for blood
sampling (36, 86). In one study, however, recordings of
blood conductivity during bolus injections of 5% glucose were used to
successfully estimate cardiac output in mice as well as rats
(86). Another alternative is the use of pulsed-Doppler
flowmetry, which has been used both invasively and noninvasively in
mice (41, 68). In our studies, a 1.5-Fr. Doppler flow
probe with a 20-MHz crystal was used in conjunction with a Doppler
flowmeter from Millar instruments. Surgical preparation is nearly
identical to that described above for the Millar Mikro-Tip pressure
transducer, except that the tip of the Doppler crystal is positioned in
the ascending aorta ~5 mm from the aortic valve and guided by the
online recording so that a maximum peak flow velocity and stable wave
form are achieved. This technique has the advantage of being able to
monitor online changes in pulsatile blood flow but the disadvantage of only measuring flow velocity, which is difficult to convert to absolute
measurements of bulk flow.
Perivascular flow probes have also been effectively employed in mice,
and although their use was originally limited by the size of the
available probes, more recent models have been miniaturized sufficiently to permit faithful recordings of regional blood flow and
cardiac output. Transit-time flowmetry from Transonics Systems (Ithaca,
NY) has become the gold standard for measuring blood flow in the mouse
and has been applied to measurements of cardiac output in the
open-chest preparation (26), as well as renal (33), descending aorta (31), and carotid
(21) blood flows. The transit time flow probes are
generally simple to use and very reliable and require surgical
isolation of only a short segment of artery (2-4 mm). After the
probe is placed around the desired artery, the acoustical path is
filled with a couplant. We easily manage to measure renal, carotid,
hindlimb, and mesenteric blood flows in mice as small as 20 g. A
new generation of probes is also available that should more readily
permit chronic placement of probes for monitoring blood flow in the
awake animal. This advancement is especially welcome in terms of
measuring cardiac output, because various methods have yielded vastly
contrasting values. Open-chest measurements of cardiac output using
flow probes generally yield values of 3-6 ml/min, whereas
preliminary measurements in closed-chest animals are in the area of 15 ml/min. This latter value would be more consistent with other
techniques used to evaluate aortic flow, including conductivity
dilution (~15 ml/min) and radioactive microspheres (~16 ml/min)
(1, 86).
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RENAL MEASUREMENTS |
The techniques applied for the evaluation of renal function in
mice have been previously reviewed and are similar to those used in the
rat, except of course for limitations in the size of plasma and urine
samples (56, 63, 69). Analysis of pressure and flow
relationships in the kidney present the same challenges as in
cardiovascular studies as discussed above. Here we will discuss three
major approaches for measuring renal function: balance studies in awake
mice, clearance and hemodynamic studies in anesthetized mice, and
micropuncture studies.
Balance studies.
Analysis of long-term electrolyte balance has been successfully
performed in mice, using either rat metabolic cages or specially designed cages for mice, which are now available commercially. In our
lab we use a simple cage design consisting of a Plexiglas cylinder,
divided into an upper chamber, which houses the mouse, and a lower
chamber, which houses a tear drop-shaped glass ball that effectively
separates feces from urine (55, 64, 74). Mattson and
coworkers (11, 62) combined balance studies with the
implantation of indwelling arterial and venous catheters to permit
monitoring of blood pressure, blood sampling (with simultaneous replacement), and electrolyte infusion. It is essential to recognize that the blood volume of a typical 30-g mouse is very small, perhaps 2-2.5 ml, and therefore that blood sampling must be kept to a minimum. A blood sample of even 100 µl represents a significant hemorrhage and can be expected to have hemodynamic consequences. When
possible, blood samples should therefore be replaced with an equal
amount of blood from a donor animal.
We managed in our lab to miniaturize many of the common assays for
evaluation of renal function to the point where only a few microliters
of sample are required. New generation blood gas instruments are able
to analyze several electrolytes on small capillary samples. For
example, we use a Chiron model 348 blood gas analyzer (Medfield, MA)
that can measure PO2,
PCO2, pH, Na+, K+, and
Cl (or Ca2+) on 40 µl of whole blood. Using
a Corning model 480 flame photometer (Medfield, MA), we are also able
to measure Na+ and K+ on only 4 µl of plasma
or urine by manually diluting samples rather than using the automatic
diluter. Plasma and urine osmolality can be measured using a Fiske
One-Ten freezing-point depression osmometer (Norwood, MA), and the
15-µl sample can be largely recovered after the analysis. A Labconco
model 442 digital chloridometer (Kansas City, MO) can measure
Cl on 10-µl samples. Finally, we used a standard picric
acid-based creatinine assay to measure endogenous creatinine levels in
mice (available from Sigma, St. Louis, MO; procedure No. 555). We
miniaturized this assay by using half-area 96-well microplates so that
the total reaction volume is 120 µl. However, creatinine
concentration in mouse plasma is very low (~0.25 mg/dl),
necessitating the use of rather large sample volumes in the assay: we
typically use 40 µl of plasma and 80 µl of alkaline picrate to
perform the assay. It is also important to note that mouse plasma
contains a large amount of non-creatinine cromagens; for this reason it
is imperative to measure the absorbance of the sample before and after
acidification of the reaction volume, the difference being equivalent
to the level of creatinine-picrate complex. Blood samples can be
obtained from awake mice by saphenous vein puncture or by tail cut and in sedated mice by retroorbital puncture (43).
Clearance and hemodynamic studies.
In acutely instrumented mice, measurements of electrolyte excretion,
inulin clearance, and para-aminohippurate (PAH) clearance are commonly used. To evaluate glomerular filtration rate (GFR), we
successfully adapted the use of FITC-inulin for use in both whole
kidney and micropuncture samples (57). For whole kidney clearance measurements, animals are infused with a 1% solution of
FITC-inulin (Sigma) at 0.15 µl · min1 · g body wt1.
Plasma samples (20-30 µl) are drawn midway through each urine collection period, with replacement of whole donor blood. Plasma or
urine aliquots of 4 µl are then diluted with 196 µl of 10 mM HEPES
buffer (pH 7.4) into 96-well microplates. The samples are then analyzed
using a microplate fluorometer with an excitation at 485 nm and
emission at 538 nm. In our experience, GFRs range between 0.8 and 1.0 ml · min1 · g kidney wt1
(57, 60), which is consistent with most of the current
literature (8, 9, 35, 78) and is also consistent with
values obtained from rat. It is interesting to note that on a kidney
weight basis, GFR in mice and rats are comparable, but when corrected
for body weight, GFR in the mouse is perhaps twice that in the rat,
reflecting the increased kidney weight-to-body weight ratio in mice.
The estimation of renal plasma flow using PAH has also been used in
mice by several investigators. Spurney and coworkers (78) used [3H]PAH to measure renal plasma flow and reported
values ~2.5 ml · min1 · g kidney
wt1. More recently, several groups reported the use of a
colorimetric assay for evaluating PAH clearance in mice infused with
2-5% PAH (9, 88), but the findings were limited due
to sampling restrictions. These investigators reported PAH clearances
ranging from 2 to 4.5 ml · min1 · g
kidney wt1. We miniaturized a colorimetric assay for PAH
(90), enabling measurement on as little as 10 µl of
plasma. In this assay we dilute plasma and urine sample 1:10 with
dichloracetic acid and, after centrifuging, mix 40 µl of the
supernatant with 40 µl of the color reagent
(p-dimethylaminobenzaldehyde) into 96-well microplates, which are read at a wavelength of 450 nm. By cannulating the renal vein
to sample renal venous effluent blood, we found that the extraction
ratio for PAH generally ranges between 0.8 and 0.9 at plasma PAH
concentrations up to at least 0.08 mg/ml. These values compare
favorably to those obtained in rats, which are generally reported to
range between 0.6 and 0.9. It is valuable to note that the plasma
concentration of PAH remains <0.1 mg/ml even at infusion rates of 6 µg · g body wt1 · min1
(corresponding to an infusion of 4% PAH at a rate of 3 µl/min). Values for renal blood flow obtained from these experiments averaged ~3 ml · min1 · g kidney
wt1.
Direct measurements of renal blood flow using flow probes are generally
preferable to estimates using PAH clearance, and current technology
using transit time and/or laser-Doppler flowmetry permits such
approaches. Gross and coworkers (32-35) evaluated
renal blood flow and pressure-natriuresis responses using a Transonics
Systems 0.5 mm V-series perivascular flow probe to measure total renal blood flow and two fiber optic strands in conjunction with a Transonics laser-Doppler flowmeter to determine cortical and medullary flow. They
reported total renal blood flow values of ~7
ml · min1 · g kidney wt1,
which correlates reasonably well with values obtained from PAH measurements. These investigators also found that total, as well as
cortical renal, blood flow was well autoregulated between 80 and 140 mmHg in normal mice but that medullary blood flow was not
autoregulated. These investigators used long-term changes in renal
perfusion pressure (induced by ligating the celiac and mesenteric
arteries and lower abdominal aorta) to evaluate autoregulatory behavior. We and others used an aortic clamp to transiently alter renal
perfusion pressure to evaluate autoregulation (82). Sample tracings from these experiments are shown in Fig.
6, demonstrating efficient autoregulatory
behavior in the mouse above ~90 mmHg. Although these data are
qualitatively similar to those obtained from rat, the upper end of the
pressure-flow relationship has not been determined in the mouse,
because it has proved to be difficult to increase renal perfusion
pressure above ~150 mmHg.
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Fig. 6.
A: sample traces of renal perfusion pressure
(top) and blood flow (bottom) during manual
reductions in perfusion pressure. The mesenteric, celiac, and lower
abdominal aorta were ligated to increase ambient blood pressure and an
aortic clamp was placed around the aorta just above the left renal
artery to allow for transient reductions in renal perfusion pressure.
B: resulting pressure-flow relationship demonstrating
efficient autoregulation >90 mmHg. Flow was measured using a
Transonics Systems 0.5 V series blood flow probe and T106 flowmeter.
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Micropuncture studies.
In vivo approaches for renal micropuncture and microperfusion have
proved to be quite feasible, requiring only moderate adaptation of the
techniques used in rat (see Refs. 71, 72 for
review). For example, Wang and coworkers (89) reported in
situ microperfusion of proximal tubules to evaluate net rates of fluid
and bicarbonate reabsorption in Na/H exchanger isoform 3 (NHE3)
knockout mice. Free flow micropuncture measurements were reported in
transgenic mice as early as 1994 (77) and since then,
several different mutant models have been studied by analyzing samples
obtained from both proximal and distal tubules (60, 73,
84). Free-flow measurements yielded a profile of nephron
function that is not altogether different from that seen in the rat.
Although values of single nephron GFR are lower in mice, averaging
perhaps 12-15 nl/min compared with 30-35 nl/min in rat,
fractional reabsorption from late proximal and early distal puncture
site is comparable to the rat: 40-50% and 70-80%,
respectively, in normal animals. In practical terms, we found that
proximal punctures in the mouse are on the same size and flow scale as
distal collections in the rat and mouse distal collections are somewhat
smaller. Preparation of the kidney, although similar to that in the
rat, requires considerably more care due to the small size. Also, the
ureter of the mouse kidney is tightly adherent to the medial margin of
the kidney and is therefore very difficult to dissect free without
bleeding. For this reason, we modified the Lucite kidney holder by
adding a second opening: the first opening is located traditionally, near the center of the holder, so as to accommodate the renal vessels
emerging from the hilum, and the second opening is located in one
corner so as to accommodate the ureter emerging from the caudal pole of
the kidney.
We miniaturized the technique for evaluating FITC-inulin clearance to
include measurements of single nephron GFR (57). With the
use of this approach, micropuncture samples (5 nl or greater) are
deposited between oil columns in a small 1-µl microcapillary tube
(microcaps, Drummond Scientific, Broomall, PA). After 0.5-1 nl of
500 mM HEPES (pH = 7.4) is added to each sample to normalize pH,
the microcaps are placed on the stage of an inverted microscope fluorometer (of the sort typically used for intracellular calcium measurements). With the use of an excitation wavelength of 480 nm and
emission wavelength of 530 nm, the samples are digitally imaged and
then analyzed for fluorescence intensity. This technique has the
advantages of being simple to use, highly sensitive, inexpensive, and
nonradioactive; furthermore, it can be performed on samples as small as
5 nl and it does not consume the sample.
Evaluation of tubuloglomerular feedback (TGF) responses, primarily by
measuring changes in stop-flow pressure in the open-loop configuration,
has also proved to be feasible in a wide variety of genetic mouse
models, and the technique is the same as that used in the rat. We
found, for example, that TGF responses in NHE3 knockout mice are intact
compared with their wild-type littermates (60). In these
studies, the stop-flow pressure measurements were also supported by
analysis of proximal-distal differences in single neprhon GFR, which
compares the filtration rate under conditions of intact and interrupted
flow to the macula densa and can be viewed as a measure of the
prevailing strength of the TGF signal at the time of the measurement.
NHE3 knockouts demonstrated a significant difference in distal vs.
proximal single nephron GFR, reflecting the persistence of a robust TGF
response in these animals. Sample traces of stop-flow pressure at
varying tubule perfusion rates obtained in our laboratory are shown in
Fig. 7. It is interesting to note that
the dynamic range of the TGF response is much lower than in the rat;
that is, the loop flow rate causing a half-maximum response is between
10 and 15 nl/min (~14 nl/min in the example shown), consistent with
the lower endogenous flow rate observed in the mouse. These low flow
rates are difficult to discriminate because of limitations in the
perfusion equipment, and whereas differences in maximal responses
should be apparent using the stop-flow technique, shifts in the TGF
relationship may be more difficult to discern in mice. This possibility
is perhaps illustrated in the study noted above (56).
Although stop-flow pressure measurements and proximal-distal single
nephron GFR differences both confirmed activity of the TGF system in
NHE3 knockouts, the stop-flow data suggested that the system was not different between the two groups of mice, whereas the proximal-distal data suggested that the strength of the TGF signal was augmented in the
knockouts. Similar findings have been reported in other strains of mice
(83).
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Fig. 7.
A: tracings of arterial blood pressure (top),
tubule perfusion rate (middle), and stop flow pressure
(bottom) during a micropuncture experiment. A wax block was
placed into the early proximal tubule to block flow; a perfusion
pipette connected to a nanoliter infusion pump was inserted into the
late proximal tubule to perfuse the loop of Henle and various rates,
and a pressure pipette connected to a servo-null pressure device (WPI
Micropressure System) was inserted upstream from the wax block to
measure stop-flow pressure. Loop of Henle flow rate was then altered as
shown. B: tubuloglomerular feedback curve showing
relationship between loop of Henle perfusion rate and the change in
proximal stop-flow pressure.
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PULMONARY MEASUREMENTS |
As in cardiovascular and renal investigations, analysis
of pulmonary function can be performed chronically in awake mice by whole body plethysmography or acutely in anesthetized mice by measurement of airway pressure-flow relationships. Parameters that can
be readily evaluated include breathing frequency, tidal and minute
volumes, airway reactivity (resistance), lung compliance, and diffusion capacity.
Plethysmography.
Several commercial systems are available for plethysmographic
measurements of respiratory function in mice and these can be found in
single chamber or two-chamber models. We have had experience with the
BioSystem XA from Buxco Electronics (Sharon, CT), which can
continuously evaluate a number of derived parameters in unrestrained, awake mice. This unit consists of a whole body chamber and integral reference chamber and pneumotachograph. A bias flow of various gas
mixtures (O2 and N2), controlled by needle flow
valves, can be delivered to a port on the chamber to maintain
O2 concentration within the chamber at a desired level. An
aerosol inlet port is also available for delivery of nebulized
bronchoactive agents. When this plethysmograph is sealed, a respiring
mouse creates pressure fluctuations, which relate to the animal's
tidal volume when the animal is breathing quietly and to the effort
of breathing when the breathing is labored. Alternatively, when the
pneumotach is open, fluctuations relate to the animal's flow rate when
the animal is breathing quietly and to the effort of breathing when the
breathing is labored. Analog pressure/flow signals from the plethysmograph are analyzed by the software package to evaluate the
following parameters: inspiratory and expiratory time, peak inspiratory
and expiratory flow, tidal volume, relaxation time, minute ventilation,
frequency of breathing rate, end-inspiratory and end-expiratory pause,
and enhanced pause (Penh). Use and validation of this
system for measuring these variables was previously published (39) and the technique has been used extensively.
To evaluate the effectiveness of this system for monitoring alterations
in ventilation in a normal mouse, we measured the above parameters
under normal conditions and immediately after exposure to hypoxia (21%
and 10% O2, respectively). As the data in Fig.
8 demonstrate, a distinct
hyperventilation (increases in breathing frequency, tidal volume and
minute ventilation) in response to breathing low O2 is
evident using this system. Investigators should be aware, however, that
these types of measurements must be made in resting mice that have been
well acclimated to plethysmograph chamber. We observed that active
(nonresting) mice typically display irregular,
high-frequency/low-volume breathing patterns (as high as 650 breaths/min), which are consistent with virtually constant sniffing
behavior.
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Fig. 8.
Sample tracings from a Buxco plethysmograph showing airflow into
and out of the chamber (Box Flow) during quiet breathing under normal
oxygen conditions (A: 21% O2) and during
hyperventilation with hypoxia (B: 10% O2).
Calculated variables from the Buxco BioSystem XA during normoxia and
hypoxia are shown in C. Ti, inspiratory time; Te, expiratory
time; PIF, peak inspiratory flow; PEF, peak expiratory flow; TV, tidal
volume; f, breathing frequency; MV, minute ventilation; RT, relaxation
time; EIP, end-inspiratory pause; EEP, end-expiratory pause; Penh,
enhanced pause.
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Airway responsiveness to bronchoconstrictor challenge is probably the
most widely measured parameter using whole body plethysmography. Generally, increasing doses of methacholine, ranging from 1 to 100 mg/ml, are delivered to the animal by aerosol through a port in the top
of the chamber. Although airway resistance cannot be directly measured
using a single chamber plethysmograph, one of the derived
parameters, Penh, has been shown to reflect changes in
airway resistance under appropriate conditions. This dimensionless parameter is based on the notion that during bronchoconstriction, the
main alteration in the pressure signal from the chamber occurs during
early expiration (when the air in the lungs become more greatly
compressed) and that the resulting change in the recorded waveform can
be quantified by comparing the chamber pressure during early expiration
with the pressure during late expiration. Because the calculation of
Penh is related to respiratory timing, rather than on
pressure-flow relationships, it is perhaps most effectively used as a
screening tool for evaluating differences in airway reactivity between
groups of mice. It is recommended, therefore, that any observed
differences in Penh be confirmed by a more thorough analysis of pressure-flow relationships to evaluate airway resistance.
Pulmonary mechanics and airway reactivity.
Whole body plethysmography provides a convenient noninvasive and
effective method for screening pulmonary function over extended periods
of time. However, to more precisely evaluate pulmonary mechanics and
airway reactivity, acute methods can be employed to generate a variety
of functional parameters in the mouse. Again there are several
commercial systems available that offer the necessary instrumentation
and software to make sophisticated measurements of lung function. Such
systems generally involve intubation or surgical tracheotomy and
subsequent measureme
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