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The Working Heart
Marc W. Merx and Jürgen Schrader
Introduction
Since the advent of genetically altered mice, a vast amount of fascinating cardiovascular phenotypes have been described with even more remaining to be explored
(Doevendans et al. 1998; Gödecke and Schrader 2000). In addition to the many ways
of assessing cardiac function in the intact animal through acute or chronic instrumentation or even non-invasively by echocardiography or MRI, the isolated, perfused
heart remains a prominent investigative tool. This is because it offers a whole array of
unique advantages:
The isolated heart provides a highly reproducible preparation that can be studied
in a time and cost efficient manner. It allows a broad spectrum of biochemical, physiological, morphological and pharmacological indices to be measured, permitting detailed analysis of intrinsic ventricular mechanics, metabolism and coronary vascular
responses. These measurements can be obtained without the confinements of systemic interference and side effects encountered in whole animal studies such as sympathetic and vagal stimulation, circulating neurohormonal factors and changes in
substrate supply as well as alterations in systemic and pulmonary vascular resistance
and left and right ventricular loading. Furthermore, experimental conditions such
as ventricular pre- and after-load, perfusate oxygenation, substrate supply, coronary
flow, heart rate and temperature, to name but a few, can be altered with ease and great
precision to address the experimental question of interest. In addition, the measurement of physiological parameters is facilitated by the convenient exposure of the isolated heart, as is the application of pharmacological agents directly into the coronary
circulation. The latter aspect also makes the working heart ideally suited for metabolic
studies. Labelled precursors can be readily applied via the coronary system and their
fate within the heart can be studied in a time-dependent manner. The preparation
also readily allows the induction of whole heart or regional ischemia at various degrees of flow and at various degrees of oxygen deprivation. Thus, the isolated heart
preparation is amenable to reperfusion or reoxygenation at various rates and with
various reperfusate compositions, providing a powerful tool for assessing many aspects of ischemia- and reperfusion-induced injury.
The isolated, working heart preparation established under strict standards represents a functionally and metabolically stable system well suited for several hours of
studies (Decking et al., 1997). However, as an ex vivo preparation, the isolated heart
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demands the application of appropriate precautions to maintain the organ’s stability
(see below). In addition, rigorous testing of the preparation must be performed and
organs failing to live up to previously determined standards should be excluded from
experimental analysis. These procedures should be repeated intermittently throughout the experimental course.
In principle two different isolated heart models exist:
▬ the isolated heart according to Langendorff (1895), in which hearts are supplied
with coronary flow through retrograde perfusion (described in the previous chapter) and
▬ the working, fluid-ejecting heart, in which hearts are perfused via the left atrium
and eject fluid through the left ventricle into the aorta thus perfusing their own
coronaries.
The latter method was first described by Neely (Neely et al. 1967) and has since been
adapted to a multitude of species including mice (Bardenheuer and Schrader 1986;
Ng et al. 1991; Grupp et al. 1993; De Windt et al. 1999).
The working, fluid-ejecting heart performs pressure-volume work, an important
distinction from its Langendorff counterpart, which performs energetically less demanding isovolumetric contractions. Because the left ventricle is filled with perfusate,
left ventricular pressure and its derivative parameters can be obtained directly through
the application of a pressure transducer. This eliminates the need for a balloon to be
inserted into the left ventricle and thus avoids the confinements inherent to balloons
such as: differences between ventricle and balloon geometry, compliance issues of
balloon material, signal dampening due to minute air inclusions in the balloon system
and the like.
With the working heart ejecting fluid, cardiac output, being the sum of aortic and
coronary flow, can be measured continuously. Preload and afterload may be adjusted
over a wide range of feasible loading conditions enabling detailed studies of ventricular function. Thus the isolated working heart adds functionally important parameters
to the already broad arsenal provided by Langendorff preparations, while retaining
ease of access to the organ and versatility in experimental design.
Description of Methods and Practical Approach
The Working Heart Apparatus
Since the first description of the working heart by Neely (Neely et al. 1967), this fascinating model has been eagerly welcomed by the scientific community and is thus
found in many laboratories throughout the world. Numerous institutions use homebuilt apparatus enabling them to add modifications according to their individual
research interests. In addition, several companies offer commercially available working heart setups (Experimetria, Hugo Sachs Elektronik/Harvard Apparatus for example). As it would be impossible to discuss all of these individual approaches to the
working heart, we will concentrate on the model employed in our laboratories
to illustrate essential components. The apparatus referred to is commercially available
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and produced by Hugo Sachs Elektronik/Harvard Apparatus. A diagram of the working heart setup and a photograph of the actual setup in our laboratories are given in
Fig. 1a and b, respectively. A schematic diagram of the work-performing heart is given
in Fig. 2a alongside a close up view of the heart in its actual experimental surrounding (Fig. 2b). An important advantage of this setup is that it facilitates seamless
switching from Langendorff to working heart mode and, while in Langendorff mode,
from constant pressure to constant flow perfusion. Furthermore, it is very compact
and can easily be transferred from one location to another as all essential components
are robustly mounted within the supportive structure. The whole apparatus is made
of Plexiglas greatly reducing the risk of accidental damage as compared to glass constructions.
The components depicted in Fig. 1 include all necessary parts for the successful
operation of a working heart model except for the thermostatic circulator and
the measuring instruments. As the isolated heart is very vulnerable to temperature
changes, every effort should be made to ensure constant and defined temperature
conditions. To this end, the heart and all temperature sensitive components are installed inside a thermostated chamber (15) and (16).
The heart is connected to the apparatus by removable aortic (2) and atrial (29)
cannulae. The cannulae can be made of glass or plastic but stainless steel is most commonly used. Cannula size is critical as, in contrast to Langendorff perfused hearts,
total cardiac output, of which coronary flow is but a fragment, flows through both
atrial and aortic cannulae. In addition, the cannulae are rigid, resulting in an energetically challenging resistance to left ventricular ejection in the case of the aortic cannula. The inner diameter of the cannula should therefore be at least the same as the
aortic diameter and preferably as large as feasible. Due to the considerable elasticity
of the aorta it is possible to stretch the aorta over a cannula with an outer diameter
significantly larger than the original aortic diameter. The length of the cannula should
be kept as short as possible for the same reasons (cannula design is discussed in
elegant detail by De Windt et al. [1999]). Even if working with one species only, it is
advisable to have several sizes of cannulae available in order to be able to employ the
largest feasible for any given heart. For our mouse studies we use cannulae of 0.9
to 1.3 mm inner diameter in 0.1 mm graduations. These cover hearts ranging from
approximately 100 mg to 600 mg wet weight and thus are applicable in mice as small
as 18 g with no relevant upper weight/size limit. The same considerations hold true
for the atrial cannula, although size constraints set by the individual heart studied
pose less of a challenge in the atrium as the pulmonary vein orifice can easily be enlarged to accommodate a sufficiently sized cannula. Nevertheless, the cannula should
present as little resistance to inflow as possible. In general, the atrial cannula should
have a free flow (i.e. flow measured without the heart being attached to the cannula)
of at least twice the flow expected to occur in the individual experimental setting. In
our experience the maximum cardiac output generated by a working mouse heart
(using high preload (25 mmHg) and low afterload (60 mmHg), see also experimental
examples section below) lies between 20 and 25 ml/min. However, these high flows are
only encountered in experimental designs where this unusual loading combination is
of interest. The atrial cannula we are able to use in all but the smallest hearts, with an
inner diameter of 1.3 mm is sufficient to permit a free flow of 70 ml/min at 12 mmHg
preload.
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2
Figure 1a
Explanation see next page, part b
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Figure 1a,b
Schematic diagram of (a) and
actual (b) working-heart apparatus. Important components
are marked and numbered as
follows (see text for detailed
explanation): 1 aortic block;
2 aortic cannula; 4 flow probe
(coronary/aortic flow); 5 flow
resistor; 6 rotary control pressure pump; 7 pressure gauge;
8 compliance chamber; 11 roller pump; 12 thermostated reservoir; 13 carbogen; 14 gassing filter; 15, 16 thermostated
chambers; 18 pressure transducer (for perfusion pressure/
afterload); 24 roller pump;
26 preload vessel; 26a suc-tion
tube; 27 flow probe (atrial
flow); 29 atrial cannula; 30
pressure transducer (preload
measurement); 32 waste reservoir; 34 dissecting microscope
11
10
15
34
16
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5
26a
4
8
26
2
27
29
y
Figure 2a
Explanation see next page,
part b
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26a
4
8
26
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y
Figure 2a,b
Schematic diagram of (a) and actual (b) working-heart preparation. Important components are marked
and numbered as follows (see text for detailed explanation): 2 aortic cannula; 4 flow probe (coronary/
aortic flow); 5 flow resistor; 8 compliance chamber; 26 preload vessel; 26a suction tube; 27 flow probe
(atrial flow); 29 atrial cannula; PE-tubing connected to pressure transducer
The perfusate is oxygenised in a thermostated reservoir (12) by gassing through
a glass filter (14) with carbogen (a gas blend of 95% oxygen and 5% carbon dioxide). Carbogen flow is regulated through a needle vent (13). If the perfusate employed is prone to foaming (see perfusate) a membrane oxygenator should be used
instead.
Although the fluid-ejecting working heart is the aim, it is nevertheless necessary
to perfuse the explanted heart in a retrograde fashion initially. Thus the apparatus is
operated in Langendorff constant pressure mode (see also atrial cannulation and
switch to working heart mode) with a roller pump (11; e.g. Reglo Digital ISM 834)
supplying perfusate to the heart. To ensure constant pressure, the above-mentioned
pump supplies more perfusate than actually required by the heart being perfused. The
excess perfusate supplied by the pump and not required by the heart passes through
a flow resistor (5) back to the reservoir (12) for oxygenation. It is thus possible to
adjust the perfusion pressure by setting the flow resistor (5) to whichever perfusion
pressure is desired, ensuring constant perfusion pressure over a wide range of flows.
The flow resistor consists of a membrane to which perfusate is channelled from one
side and pressure is applied from the other side, using a rotary control pressure pump
(6) connected to a pressure gauge (7) indicating the actual perfusion pressure. In
addition perfusion pressure is measured by a pressure transducer (18; e.g. ISOTEC
single use, comparable-pressure transducers also available from Braun, COBE or
Millar). The resulting coronary flow is measured with a flow probe (4; e.g. Transonic
1 N connected to T106, T206 or HSE-TTFM). After passing through the flow probe
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and a compliance chamber (8) doubling as bubble trap, the perfusate reaches the
aortic cannula (2). A frequently used, albeit less versatile, setup consists of a pressurized water column with the height of the water column determining perfusion pressure.
If constant-flow perfusion should be required (this is not routinely recommended
when initiating a working heart experiment), return flow of perfusate supplied by the
pump (11) is prevented by setting the control (6) of the flow resistor (5) to a very high
value (e.g. 250 mmHg) thus sealing the membrane of the flow restrictor. The required
flow is then set on the pump (11). The resulting perfusion pressure is measured with
the pressure transducer (18) and coronary flow rate is verified using the above mentioned flow probe (4).
In working-heart mode, the left atrium receives perfusate from the preload vessel
(26) through the atrial cannula (29). Using a roller pump (24; e.g. Reglo Digital ISM
834) more perfusate is pumped into the preload vessel (26) than flows into the atrium.
The preload vessel doubles as a bubble trap. The solution not required is pumped
from the stainless steel suction tube (26a) back to the reservoir using one of the pump
channels.
Preload is altered by vertical adjustment of the suction tube in much the same way
as afterload would be adjusted in the case of a setup with a pressurized water column
(although the pressure range being covered has to be much larger in the latter case;
26a). Preload is measured by a pressure transducer connected to the preload vessel
(30) (e.g. ISOTEC P75). The resulting atrial flow (equalling cardiac output in the working-heart model) is measured continuously with a flow probe (27; transonic 1 N, see
above).
The perfusate ejected by the left ventricle passes into the coronaries by way of the
aortic root. Perfusate ejected by the heart but not required for coronary perfusion
(which may be viewed as “effective cardiac output”) flows into the aortic block (1) via
the aortic cannula (2). Here it enters the compliance chamber which contains an
adjustable amount of air (we recommend 1 ml) to provide elastic recoil (simulating
the “Windkessel” characteristic of the aorta). The compliance chamber functions
as a buffer against the low mechanical compliance of the setup as such. It lowers
mechanical resistance to the ejecting heart and ensures coronary perfusion which
occurs mainly during diastole in the work-performing heart. From the compliance
chamber the perfusate runs through the adjustable flow resistance (5) that is now used
to set afterload in the same way that it was previously used to adjust perfusion pressure. Via tubing (5a) connected to the flow resistor (5) the perfusate ejected by the
heart is drained passively either to the oxygenating reservoir (12), providing recirculation of the perfusate, or alternatively to a waste reservoir (32). Afterload is now
measured with the same pressure transducer previously used to measure perfusion
pressure (18).
Aortic flow produced by the heart at any given afterload is measured continuously
via flow probe (4). Again coronary flow is assessed with the same flow probe as in
Langendorff mode. It is important to note that if the same flow probe is used for
Langendorff and working-heart measurements, polarity has to be changed when
switching from one mode to the other, as flow direction will change also (otherwise
negative flow values will result). The difference between atrial flow (i.e. cardiac out-
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put) and aortic flow constitutes coronary flow in the working heart model. Depending on the data acquisition system preferred, the latter can be computed and displayed
online (see below) or calculated after the experiment is completed. For calibration
purposes, the coronary perfusate having passed through the coronaries, right atrium and ventricle can be collected form the pulmonary artery and its volume measured.
More importantly the perfusate collected from the pulmonary artery may be used
to measure coronary venous PO2 . For this purpose the pulmonary artery is cannulated
with PE tubing and suctioned to a PO2 electrode with constant flow (as PO2 electrodes
are very sensitive to flow alterations, e.g. ZABS PO1750). The rate at which the coronary venous perfusate is suctioned should always be well below the rate of coronary
flow to avoid air being sucked into the PO2 measuring chamber. Excess perfusate from
the right ventricle is suctioned by a roller pump (24) through tubing (23) connected
to a draining cannula (22) at the bottom of the thermostated chamber (16). It is than
either recirculated to the oxygenation reservoir (12) or drained to the waste reservoir
(32).
Left ventricular pressure is measured with a fluid filled catheter inserted into the
left ventricle and connected to an additional pressure transducer (not shown in the
diagram, e.g. ISOTEC MP15). Alternatively, a bespoke catheter with miniature pressure transducer mounted to its tip can be used for even truer pressure readings albeit
at considerable cost (Millar, Radi).
Recently, an even more sophisticated (and costly) alternative has become available
in the form of pressure volume catheters (Millar). In the latter case, a miniature pressure transducer at the catheter’s tip is combined with induction coils at the catheter’s
distal end for volume measurements.
The electrocardiogram of the work-performing heart can easily be recorded by
applying electrodes to the right atrium and ventricle. It is also possible to record the
electrocardiogram via the atrial and aortic cannulae, although we have found the
signal quality to be inferior to directly attached electrodes. If pacing of the heart is
required in the experimental protocol, this can be achieved through the application of
a, preferably bipolar, stimulation-electrode connected to a stimulator suitable for the
high frequencies of up to 15 Hz required (e.g. Hugo Sachs Elektronik/Harvard Apparatus P105).
A further important consideration is the application of drugs to the heart to enable pharmacological studies. In principle, it is possible to add the agent directly
to the perfusate (see also Perfusate). However, if different concentrations are to be
studied or if several agents are of interest during the course of one experiment, it
is more convenient to add the substance through a sidearm of the apparatus with a
receptacle for one or more infusion lines. This sidearm should be as close to the atrial
cannula as possible (or to the aortic cannula while in Langendorff mode). In the setup described here a sidearm is integrated directly proximal to both the atrial and aortic cannulae, thus enabling the precise application of even minute volumes through
micro-infusion pumps (e.g. Precidor P104).
In addition to circumventing the need for several perfusate reservoirs with all
the associated drawbacks (see perfusate), microinfusion through a proximal sidearm also avoids contamination of the whole setup wit the substance studied, which
can be responsible for uncontrolled side effects due to prolonged wash out.
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With the apparatus described here, preload can be set in the range of 0–30 mmHg
and afterloads ranging from 0 to 300 mmHg are feasible. Atrial and retrograde flow
(i.e. coronary flow in Langendorff) depend largely on the size of the cannulae and
the pump and tubing employed. In our setup tailored to mice, atrial flows of up to
50 ml/min and retrograde flows of up to 25 ml/min are possible (using the cannula
sizes mentioned above). Preload, afterload and left ventricular pressure as well as
atrial (sometimes referred to as venous) and aortic (sometimes referred to as arterial)
flows are measured continuously. Depending on the recording hard and software,
a multitude of derivative parameters can be calculated online or after completion of
the experiment.
Preparatory Steps
Prior to the first experiment of the day the working heart apparatus should be flushed
with purified water (i.e. ultra-filtered water generated by e.g. a Millipore System). In
general, letting the apparatus become dry should be avoided, as it is extremely tedious
to eliminate small air bubbles adherent to the walls of the apparatus and tubing. If
meticulous attention is not paid to keeping the setup free of air, air emboli can cause
coronary occlusion in the isolated organ. It is thus advisable to keep the apparatus
filled with purified water between experiments, unless it is decommissioned for long
periods of time. Leaving the perfusate itself in place between experiments for more
than a few minutes has the drawback of promoting crystallisation on apparatus and
tubing walls. If the perfusate is left in place for hours, there is a risk of promoting
microbial growth and thus colonization of the apparatus, a condition that is extremely
difficult to treat and that may demand complete dismantling of the apparatus and
replacement of significant parts. Along the same lines, the heating (and where applicable cooling) circuit of the apparatus should be kept as clean as possible by
frequently replacing the purified water (i.e. ultra filtered water, see above) and by adding stabilizing antimicrobial agents.
Depending on the surrounding temperature it is necessary to initiate heating (and
cooling) of the apparatus well before starting the experiment, as stable temperatures
are essential to generate reproducible results. For mice the temperature of the perfusate provided at the tip of the aortic perfusion cannula should be 37 ± 0.5 °C. As the
body temperature of rats and rabbits is slightly higher, 38 ± 0.5 °C should be aimed for
in these species. The chamber surrounding the isolated heart should be heated to the
same temperature. The chamber should also be as airtight as possible to avoid loss
of humidity and minimize cooling of the isolated heart secondary to evaporation of
perfusate from its surface. If heart preparation is performed in a cooled environment
(see below) the heart should be held at a constant temperature of 4 ± 0.5 °C throughout preparation.
Perfusate
The perfusion solution should be prepared as close to the actual time of use as possible. In any case it is not recommended to store the substrate-containing perfusate for
more than a few hours, as it is an excellent bacterial growth medium and bacterial
contamination of the perfusate will thus readily occur.
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The perfusate employed in isolated heart perfusion serves two main purposes
providing (a) substrates and (b) oxygen to the heart while maintaining ionic and pH
homeostasis. Because of the associated ease of use, saline solutions gassed with carbogen (95% O2 + 5% CO2, the latter to achieve the required pH of 7.4) and with glucose
and pyruvate as substrates are employed most frequently. However, fatty acids are
used in metabolic studies and albumin has been added to the perfusate to reduce the
otherwise significant oedema associated with saline perfusion. If fatty acids or proteins are added to the perfusate, oxygenation should be carried out with membrane
oxygenators as severe foaming will result from direct gassing.
Drugs or other agents may be added directly to any perfusate or, more elegantly,
be injected through a side arm of the apparatus close to the atrial (or aortic) cannula.
If the latter technique is employed, high concentrations of the agent must be used so
that the injected volume remains minute (i.e. <1%) compared to the total flow of perfusate to the organ. Thus the amounts of substrates and oxygen delivered to the organ
remain practically unaffected. However, especially with small hearts such as those
encountered in murine studies precision pumps are necessary to apply the microliterrange volumes required (see above).
Turning to the second important function of the perfusate, the delivery of oxygen to the isolated heart, saline solutions are characterized by a relatively low oxygen
carrying capacity. This poses a challenge, as continuous provision of oxygen in quantities sufficient to maintain normal metabolism and transmembrane gradients as
well as to support aerobic ATP generation for contraction is essential for the survival
of the isolated heart. The low oxygen carrying capacity of saline solutions can be
counterbalanced by gassing with high partial pressures, as is the case with the application of carbogen.
Depending on ambient pressure O2 partial pressures in excess of 650 mmHg are
attained. Combined with the higher coronary flow observed in saline perfused hearts
compared to sanguineous perfusion, this is sufficient to ensure adequate oxygenation.
Furthermore, we were able to demonstrate that intracellular PO2 remains unaffected
even if the saline perfusate is gassed with 70% oxygen, translating into a significant
safety margin when working with carbogen (Merx 2001).
If, however, specific experimental questions require near normal coronary flow
rates and/or higher oxygen transport capacity, donor blood or erythrocyte enriched
aqueous solutions have been used successfully (Gamble et al. 1970; Qiu et al. 1993).
It should be noted that because of the volumes required in most isolated heart setups,
multiple donor animals are often required. Incompatibility of different species with
respect to erythrocytes’ ability to traverse the capillary bed of the recipient heart have
to be taken into account. Here also gassing needs to be achieved through membrane
oxygenators as direct gassing will cause excessive foaming and damage the erythrocytes and other cells in the case of whole blood perfusion. In the case of whole blood
perfusion, sufficient anticoagulation is difficult to achieve given the prolonged exposure to all kinds of non-biocompatible materials.
As mentioned above, a saline perfusion solution based on the “physiological” solution first proposed by Krebs and Henseleit (1932) is adequate for the vast majority
of isolated heart studies. The modified Krebs–Henseleit buffer used most commonly
in our laboratories contains (in mM): 116 NaCl, 4.6 KCl, 1.1 MgSO4, 24.9 Na HCO3,
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2.5 CaCl2, 1.2 KH2PO4, 8.3 glucose and 2 pyruvate equilibrated with 95% O2–5% CO2
to yield a pH of 7.4. The chemicals used should be of high purity (at least pro analysis, “p.a.” quality) and should be obtained from the same provider consistently (we
use Sigma chemicals).
As the solution contains calcium and phosphate ions, the risk of calcium phosphate particles forming and precipitating should be avoided by ensuring that the pH
of the bicarbonate buffered solution is lowered to 7.4 by gassing with carbogen and
adding calcium as the last component thereafter. Finally, it is of great importance to
filter the perfusate carefully (filter pore size 5 µm or smaller) as even the purest grade
chemicals may contain enough debris to cause embolization especially in a vasculature as small as the murine.
Heart Explantation and Aortic Cannulation
Deep anaesthesia has to be induced in the animal prior to initiation of the required
surgical procedure. In order to achieve optimal anaesthetic results with reproducible
dosing, every effort should be made to minimize stress by keeping the animal in a
quiet environment that the animal is accustomed to and by minimizing animal handling.
To prevent blood clotting in the excised heart, the animal should be sufficiently
anticoagulated prior to the application of the anaesthetic agent. This is easily achieved
by giving heparin by intravenous or intraperitoneal injection at a dose of 5 IU/g BW
(because of its lipolytic properties heparin should be avoided in lipid or fatty acid
metabolism studies). Anaesthesia is usually induced either by inhalation or by intravenous or intraperitoneal injection. Inhalative agents used in mice include ether,
halothane, isoflurane, enflurane and methoxyflurane. Intravenous/intraperitoneal
agents reported in mice include urethane, pentobarbitone, ketamine and propofol.
Relaxing agents such as benzodiazepines and curonium-derivatives as well as analgetic substances such as morphine may be added.
The anaesthetic regime will depend largely on the purpose of the study and thus
on considerations regarding the differing degree of cardio-depressant effects observed for the agents. In our labs we most commonly use a combination of ketamine
(60 mg/kg body weight) and xylazine (10 mg/kg body weight). Anaesthesia will also
be influenced by local animal welfare regulations. As substantial strain differences
(and especially in mice and rats even differences between animals of the same strain
but from different animal facilities) exist in the susceptibility to different anaesthetic
agents, careful dose titration in preliminary experiments is mandatory.
After the animal is anaesthetized, the peritoneal aspect of the diaphragm is visualized by a transabdominal incision. While supporting the sternal apex the diaphragm
is carefully incised so as not to damage any intrathoracic structure. While continuously supporting the sternal apex upward, the thorax is opened by bilateral incision
at its dorsolateral edge. A sharp but sturdy (the ribs have to be cut in the process) pair
of curved and blunt tipped scissors is usually well suited to perform the incision with
minimal risk of damage to the heart or lungs. The now mobilized anterior half of the
thoracic cage is subsequently deflected above the animal’s head to give unhindered
approach to the heart and lung. In larger species, the aorta, pulmonary vessels and
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cava can be cut directly, with some investigators even opting to cannulate the aorta in
situ. Generally speaking, the procedure can be performed rapidly (well below 1 min)
in larger species and does not pose a major challenge.
However, if mice are the species of interest it is more practical to extract the whole
heart lung package out of the thorax. The most convenient approach in doing so is
to grip the descending aorta with a pair of small forceps and cut the aorta below.
Never letting go of the aorta, the vessel is deflected ventrally and the heart and lung
dissected out of the thoracic cavity by a series of small incisions strictly parallel and
as close as possible to the dorsal thoracic wall.
If the incisions are followed through cranially the heart and lung will be removed
from the thoracic cavity by the forceps still holding the descending aorta. Heart and
lung are immediately transferred to a bath containing cold, heparinised (5 IU of heparin/ml), modified Krebs–Henseleit solution (4°C, see above) and placed beneath a
dissecting microscope.
Now the lungs, thymus, bronchi and oesophagus are dissected away taking great
care to have full visualization of the intended cutting plane to avoid involuntary
laceration of any cardiac structure. As soon as the aorta and its cranial branches are
clearly visualized, the aorta is cut just below its first branch. The heart is then gently
slipped on to the perfusion fluid-filled aortic cannula avoiding excessive strain and
thus tearing of this delicate structure. A ligature is placed as above and correct location and sufficient tightness verified under the microscope.
The cannulated heart is transferred to the apparatus. The aortic connector for the
cannula should be gently dripping with perfusate and the cannula approximated to
the connector at an oblique angle to avoid air emboli at the time of heart attachment
to the apparatus. After some practice it should be possible to perform the preparatory
steps described above in less than 3 min.
Atrial Cannulation and Switch to Working-Heart Mode
With the heart now adequately perfused in the Langendorff mode and thus oxygenated, the further preparatory steps can be taken under a less rigid time constraint.
Again a dissecting microscope (34) should be placed in front of the heart to facilitate
clear visualization of the relevant structures (see Fig. 1b, in the case of species larger than mice the naked eye should suffice, but magnification is very helpful in training).
First any surplus tissue (thymus, fat, lung ...) is removed. To ensure unimpeded
drainage of coronary venous perfusate the pulmonary artery should be incised. This
is advisable as the close proximity of the pulmonary artery to the aorta and the connective tissue surrounding both vessels result in relatively frequent, inadvertent ligation or at least partial obstruction of the pulmonary artery. The incision also
facilitates later cannulation of the pulmonary artery if arterial-venous differences in
perfusate oxygen content are to be measured. Hereafter, the dorsal aspect of the left
atrium is trimmed from surplus tissue, especially the pulmonary veins, taking care to
leave the actual dorsal atrial wall intact. In mice two pulmonary veins drain into the
left atrium, separated by a small trabecular structure. The latter is cut and the resulting combined orifice used for atrial cannulation. The atrial cannula should be drip-
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ping with perfusate when being inserted so that any air present in the left atrium or
ventricle is removed. The cannula is then tied into the left atrial wall by securing the
left atrial tissue surrounding the orifice above the flange at the distal end of the cannula with a suture.
If required, the pulmonary artery is cannulated next (i.e. to measure coronary
venous PO2). Electrodes for ECG recording or pacing are placed if desired. After instrumentation of the heart is completed (usually within 5 min after initiation of Langendorff perfusion, the heart is left for 20 min to equilibrate under retrograde perfusion
(i.e. Langendorff mode, chapter 2.1) preferably under 100 mmHg constant perfusion
pressure. It should be noted that there is no flow to the atrial cannula at this stage.
Thereafter, perfusion pressure is lowered to 60 mmHg and preload (i.e. atrial perfusion pressure) is set to 12 mmHg. After enabling flow through the aortic cannula,
flow in the aorta should be reversed as the heart begins ejecting perfusate in an antegrade fashion and the aortic perfusion pressure becomes afterload. It is sometimes
helpful to lower afterload further (to values as low as 20 mmHg) to facilitate the initiation of antegrade flow, but this should not be done for no more than a few seconds
as afterload equals coronary perfusion pressure and cardiac hypoxia would result.
After-load should then be increased to values between 80 and 100 mmHg and the
heart left to equilibrate for another 10 min, after which the experimental protocol of
choice may be started.
Data Acquisition
Afterload, preload, left ventricular pressure (LVP), atrial flow, aortic flow and if
desired coronary venous PO2 as well as the electrocardiogram are measured continuously. The signals originating from the pressure transducers and flow meters
as well as electrocardiogram and PO2 signals require amplification. This is usually
performed by tailor-made amplifiers designed specifically for the signal in question that are often bundled with the respective probe or transducer. In our laboratories we use dedicated amplifiers by Hugo Sachs Elektronik/Harvard Apparatus
(TAM-A for the pressure transducers, TTFM for the flow probes, ECGA for electrocardiogram and OPPM for coronary venous PO2) slotted into a convenient single case
(PLUGSYS).
Traditionally the amplifier output is plotted on a polygraph which allows the user
to monitor and record the parameters studied. However, this implies often tedious
manual evaluation after completion of the experiment. Analogue-digital converters
connected to personal computers running specially designed software have the advantage of calculating derivative parameters online and displaying them together with
the primary data in almost any fashion that suites the investigator.
In our laboratories we use an IBM-compatible personal computer with analoguedigital converter (Texas Instruments, 2000 Hz resolution) and specifically designed
software (EMKA Technologies, Paris, France). The above-mentioned parameters as
well as derivative parameters are displayed in real time and streamed onto hard disk
(at a rate of approx. 30 Mb per hour). The derivative parameters we work with are:
a) coronary flow, being the difference between atrial and aortic flow;
b) the maximum rate of left ventricular pressure development (dP/dtmax);
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c) the maximum rate of relaxation (also referred to as minimum rate of pressure
development; dP/dtmin)
d) time to peak pressure (TPP), calculated as time from end diastolic pressure (EDP)
to systolic pressure (SP);
e) relaxation halftime (RT1/2), time from SP to DP/2).
In-Vitro Techniques
B through e are all derived from left ventricular pressure curve analysis.
In addition to facilitating convenient display (we use a twin screen setup enabling
us to monitor the real time signal curves as well as giving us a complete overview of
all signal trends at any time) and storage of data, the software allows us to predefine
or manually select time points in the course of the experiment during which data is
written into an Excel table (Microsoft) in addition to and independent of the raw data
being streamed to the hard disk. The fashion in which the data is written to the Excel
table is freely chosen by the investigator e.g. coronary flow might be stored every 60 s
with the value stored being the mean calculated over 30 s. In this way, through thoughtful design of the experiment and of the software storage functions, a large part of
experiment evaluation can be automated.
Examples
The examples given here are taken from working heart experiments performed to
analyse differences in cardiac function between myoglobin (Mb) knockout mice
(myo–/–) and their wild type counterparts. Weights of excised hearts ranged from
180–250 mg, with no significant differences between WT and myo–/–. Left ventricular mass (including septum) as a percentage of whole heart wet weight was similar
(64.7 ± 2.1% vs. 65.3 ± 2.7; WT vs. myo–/–; p = n.s.).
Afterload was increased stepwise to simulate growing systemic resistance. As
listed in Table 1, impaired contractility was demonstrated by slower contraction and
relaxation reflected as time to peak pressure (TPP) and relaxation half-time
(RT1/2) in myo–/– hearts. As a further sign of decreased contractility, the rate of pressure development (dP/dtmax) was slower in myo–/– hearts at all afterloads studied.
Intrinsic heart rate (HR), left ventricular developed pressure (LVDP) and pressure
volume work (cardiac power) observed at the different degrees of afterload (pressure
load) did not differ.
Figure 3a depicts the parallel coronary flow pattern observed in both groups at
increasing afterloads. However, coronary flow in myo–/– hearts was almost twice as fast
as that in WT hearts at the corresponding afterloads. As cardiac output (CO) was not
increased in myo–/– hearts, resulting aortic flow was considerably smaller, even approaching zero at the highest afterload employed (120 mmHg).
Stepwise increase in preload was performed to analyze the influence of filling pressure on the given mouse hearts. Intrinsic HR, peak systolic left ventricular pressures
(LVP) and pressure volume work (cardiac power) observed at the different degrees
of preload (volume load) did not differ and rose only slightly with growing preload .
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Table 1
Cardiac parameters of myo–/– and WT controls at selected afterloads
Afterload
50 mmHg
70 mmHg
90 mmHg
120 mmHg
Heart rate, bpm
470 ± 27
481 ± 21
491 ± 17
503 ± 18
LVDP, mmHg
72 ± 2.8
93 ± 2.9
113 ± 3.8
142 ± 6.4
Cardiac power,
mmHgxml/min
2499 ± 361
3313 ± 545
3892 ± 754
4396 ± 630
dP/dtmax, mmHg/sec
4618 ± 268
5359 ± 168
6524 ± 508
7542 ± 535
TPP/mmHg,
ms/mmHg
0.38 ± 0.02
0.32 ± 0.01
0.25 ± 0.01
0.16 ± 0.01
RT1/2/mmHg,
ms/mmHg
0.42 ± 0.01
0.34 ± 0.01
0.28 ± 0.02
0.20 ± 0.02
Heart rate, bpm
473 ± 21
479 ± 20
486 ± 22
507 ± 27
LVDP, mmHg
73 ± 3.4
94 ± 3.0
115 ± 3.6
139 ± 7.1
Cardiac power,
mmHgxml/min
2628 ± 465
3358 ± 573
3882 ± 584
4467 ± 557
dP/dtmax, mmHg/sec
3883 ± 351b
4789 ± 526b
5782 ± 640b
7243 ± 733a
TPP/mmHg,
ms/mmHg
0.49 ± 0.02c
0.37 ± 0.02c
0.31 ± 0.02c
0.23 ± 0.02c
RT1/2/mmHg,
ms/mmHg
0.52 ± 0.02c
0.41 ± 0.01c
0.33 ± 0.02c
0.27 ± 0.02c
WT
myo-/-
Coronary flow remained stable at the various preloads studied (Fig. 3b) while being
approximately 55% higher in myo–/– hearts compared to their WT counterparts, i.e.
20.2±2.0 mlxmin–1xg–1 vs. 13.0 ± 1.9 mlxmin–1xg–1 (p<0.001) at 5 mmHg preload. With
CO being identical in both groups, aortic flow was significantly smaller in myo–/–
hearts compared to WT hearts, i.e. 14.7±3.3 mlxmin–1xg–1 vs. 23.4 ± 5.7 mlxmin–1xg–1
(p<0.005) at 5 mmHg preload. The lower aortic flow values found for myo–/– hearts
reflect the larger fraction of CO required to perfuse the coronaries of myo–/– hearts
adequately (Fig. 3b).
Despite the absence of significant differences in LVDP and cardiac power between
the two groups, a marked decrease in contractility was detected in myo–/– hearts at all
preloads studied. This contractile impairment was especially pronounced at low
preloads.
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In-Vitro Techniques
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Isolated Organs
Figure 3a,b
Coronary and aortic flow per gram heart weight in the working heart setup for myo–/– and WT mice
under a wide range of preloads (a) and afterloads (b). Cardiac output is calculated as sum of aortic and
coronary flow (*p<0.05; **p<0.005; ***p<0.001). For coloured version see appendix
For example dP/dtmax at 5 mmHg preload was 4688 ± 425 mmHg/sec in myo–/–
hearts versus 5718 ± 435 mmHg/sec in wild type hearts (p<0.001) while at 10 mmHg
preload the difference in dP/dtmax between the respective hearts narrowed to
6001 ± 476 mmHg/sec vs. 6447 ± 547 mmHg/sec (p<0.05).
Troubleshooting
In general, meticulous attention should be paid to keeping every aspect of the apparatus clean and well calibrated. In addition, tubing should be replaced frequently.
Although this is very time consuming and, in the case of tubing, costly, it should be
kept in mind that contamination of the apparatus or a failure to calibrate the transducers, flow meters and other sensors used can easily ruin a whole experimental
series. Furthermore, the complexity of the apparatus, connected electronics and software require the investigator to be as familiar as possible with his individual setup in
order to gain the maximum information from any one experiment. Being familiar
with the apparatus also allows the investigator to respond quickly and professionally
to most problems that might occur during the experimental course and thus save an
experiment that otherwise might have been lost.
▬ In our experience, the most common problems arising when using the working
heart originate from small air bubbles. These can be entrapped in flow probes
leading to a sudden interruption in flow measurement. Often these small bubbles
will not pass through the flow probe and may have to be flushed out. While this
The Working Heart
189
manoeuvre is readily performed prior to the organ being hung in the apparatus,
it can be a great nuisance during the experiment. Air bubbles can also significantly
dampen the pressure signal if they find their way into PE catheters used for pressure measurements or even into the transducer domes. Because even minute air
bubbles, which are extremely difficult to detect, can cause dP/dt values to fall by
30%, the whole apparatus should always be checked carefully for air bubbles and
preferably not be allowed to become dry (see also preparatory steps).
▬ The isolated murine heart is very temperature sensitive. Drops in temperature due
to opening of the heated chamber surrounding the heart, lead to a prompt decline
in function and should be avoided whenever possible during the experiment.
▬ The chemicals from which the perfusate is made should be checked on a regular
basis and they should be obtained from the same source whenever possible to
eliminate alterations from delivery to delivery. The perfusate pH should always be
controlled, as minute differences in pH greatly affect coronary flow and thus organ
performance. In this context it should be noted that if working with perfusates
being gassed by different gas mixtures, precisely 5% carbon dioxide should always
be included in the mixture to ensure a pH of 7.4. In our experience this is only
feasible through special, made to order, gas or by applying precision mass flow
controllers.
▬ Finally, the animals studied should be in good health and rigorous monitoring
routines should be in place at the local animal facility. This is especially important,
as many murine infections are not accompanied by overt clinical symptoms.
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