Q&A Report: Measurement of Cardiac Function Using Pressure-Volume Loops in Swine

Pedro Ferreira, PhD
Postdoctoral Researcher
Physiology and Cardiothoracic Surgery
University of Porto

The catheters measure conductance, or in simpler terms how easy electricity passes through the surrounding tissues (in this case blood, muscle, etc). Since blood is a better conductor then the surrounding structures, when the heart fills up with blood (surrounding the catheter with more blood), the conductance signal increases, and vice-versa. After some calibration steps (namely parallel conductance and field inhomogeneity), you can calibrate this signal into volumes and have your PV Loop. Take a look at one of the most important papers about this topic: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2597499/

We use a Ventri Cath as many times as we can! There are to ways that we decide to stop using it, either the pastic itself is very damaged (or kinked) and we are not able to place it in the correct position, or the sensors stop working. If the pressure sensor is damaged, you will likely see this by either a big drift (pressure changing significantly when the catheter is steady in the same position – for example inside a syringe with warm saline), or if the pressure signal becomes very noisy.
Alternatively, the conductance electrodes can also stop working, and you can test this by placing the catheter inside a good conductor (like saline or hypertonic saline) and see if the conductance values change in an expected way.

So far, we’ve used our catheters at least more than 20 times, and the most common reason to change is physical damage to the catheter body, which makes it kinked and hard to place in the correct position. Also, after a while, these kinks will end up damaging the inner wires and you will start losing segments.

There are some catheters which seem to be appropriate for rabits. Please see publication: https://academic.oup.com/icvts/article/26/4/673/4683722

In that publication they mention a 3Fr multisegment catheter, which according to Millar’s webpage I guess it corresponds to the SPR-877. They do this using an open chest approach in rabits with ~3.2 kg.

The tip of that catheter is straight, so I guess you could use the carotid approach to introduce it into the LV (similar to the one used for mice and rabits). I have however seen some labs doing right heart catheterization using a closed chest approach in rabits (by introducing a pediatric swan ganz), so in theory you could perform “percutaneous” LV PV Loop measurements in rabits by using a surgical approach to access the vessels, as I don’t think that it will be easy to get introducer sheaths in the arteries and veins of these rabbits. For the IVC occlusion there are plenty of options of small balloons.

Ventri Cath is a pressure volume catheter, which measures both signals, while the Mikro-Cath is a high fidelity pressure sensor that only measures pressures.

The Ventri Cath is conductance technology, which is sold by Millar / ADInstruments. It gives you conductance signals which can be calibrated to volume and pressures.

Yes. If we use a SG catheter with a thermistor (a “CCO” catheter), it can interfere with the electrodes closest to it, especially in smaller animals. It is a very clearly observed interference in the conductance signal, which stops once you stop the CO measurement. When we use these catheters we measure CO up until we are ready to make our acquisitions, stop it, acquire our data, and turn it on again to confirm hemodynamic stability. We have not see interference between the LV and the RV catheters, although we work with pigs weighing between 40 – 90 kg, I don’t know if this would happen in smaller animals.

Not that I have noticed. Most of our “moving around” is due to breathing, so if you stop ventilation you should be able to see a very stable loop. If not, probably the catheter is too far in and touching the wall or touching a papillary muscle which would ultimately cause arrythmias. We have not noticed the catheter being moved by the aortic valve. I think that the curves the catheter makes until it reaches the LV, and the places where it touches the wall give it sufficient support to keep stable.
The catheter will move around with the heart, but if the pigtail part is nicely “snug” in the apex, it will be very stable.

Most of times, at least in our experience, partial occlusion of the IVC is not sufficient to result in a fast (few seconds) decrease in preload to a sufficiently low level to obtain a good PV relationship. Furthermore, one would have to estimate how much volume to inject in the IVC balloon, or estimate the size first to determine what the partial occlusion will correspond to. If the animal is hemodynamically stable, a few seconds complete occlusion of the IVC will be quickly recovered (at least in terms of ventricular function).

It depends on the project you are working on. If we are working with a drug aimed at improving diastolic function, we mainly look at EDP, dp/dt min, tau and EDPVR, if we are looking at a drug that is aimed at improving heart failure in general, we also look at EF, CO, ESPVR, etc. We also analyze ventricular arterial coupling very often, as it can be very informative, and this is the ratio between Ees (ventricular elastance) and Ea (arterial elastance).

But pretty much most of the parameters resulting from the PV table in LabChart can be informative one way or another. My advice is to look at as many parameters as you can, since if more than 1 parameter corroborate your finding you have a lot more confidence in it.

For large animals, I am not aware of other alternatives. For small animals, I’ve seen some references to stopping ventilation during inspiration, which increases intrathoracic pressure and reduces preload, but it also increased afterload on the RV, so it is not perfect. Other researchers sometimes put abdominal pressure on top of the IVC (its relative position), but this will also put pressure on the aorta.

The only “clean” way of reducing preload, in my opinion, is the IVC Balloon. If you use a conservative approach, this is not destructive and you can recover the animal afterwards.

There is also the option of determining the ESPVR relationship using the single beat method. This would avoid using an IVC Balloon, but would have to be validated in your specific context.

No, I mean the actual loop itself having negative volumes, not the intercept. If you overestimate your parallel conductance, you can get negative volumes either already at baseline, or after performing an IVC occlusion. This is obviously wrong, so the hypertonic saline needs to be repeated. This is why I emphasized the need to always check Vp when performing the hypertonic saline administrations.

Indeed sometimes the ESPVR can assume a non linear relationship which could cause a negative V0 intercept if the linear equation is used. Assuming a quadratic function might solve that issue. Alternatively, and going back to the first point, if Vp is overestimated, even if the Loop volumes don’t become negative, V0 can be negative. This could affect parameters that are derived from the PV Area such as cardiac efficiency.

After data collection there is very little that can be done. If you assume that the animals in the same groups are very similar and that Vp is more or less stable, one could in theory apply the group value, but this is not the best approach.

Alternatively, if you have an external source of volume measurement that gives you either EDV and ESV or EF, you can confirm if the Vp is correct.

Check your Vp value to make sure that you are not overestimating it. Make sure that your ESPVR does not follow a non-linear relation.

In large animals we always record all our segments, and we create a new Channel were we sum the different segments (mimicking the composite signal).

When we analyze our data, we add up the segments that are correct (move in the counter clockwise direction), and once we’ve decided which segments to use, we use this “Volume Sum Channel” as our input for the volume signal in the LabChart analysis module. After that we follow the same principles as in the analysis for small animals.

We do not use the Rho Cuvette, as we always have an external source of CO. You could potentially use it to have a second technique to calibrate volumes and have a plan B in case something fails, or just to confirm that your Rho Cuvette and external CO match.

After we apply our Vp, we apply our alpha based on our external CO measurement. Either the thermodilution Swan-Ganz (which we do immediately before our acquisitions – 3 consecutive and consistent measurements), or the CCO Swan-Ganz (which we turn off before acquisitions), or a ultrasonic flow probe around the ascending aorta if we are working open chest.

Yes, similarly to the LV, you can introduce a sheath into the RV and place a Ventri Cath there. Our preferred approach is: 1) advance a Swan-Ganz with a large enough lumen for a 0.035 in guidewire; 2) leave the guidewire in the distal PA; 3) Advance your sheath over the guidewire into the PA; 4) Advance your Ventri Cath through the sheath, which is still in the PA, but do not push it out of the sheath yet. If you allow the pigtail to form, when you pull it back into the RV you might damage the PA valve. 5) slightly pull back the sheath until you are in the RV outflow tract (use contrast to help if needed, or follow the pressure of the sheath or the Ventri Cath), and advance the Ventri Cath, making sure it does not go into the PA.

Usually the best loop is when the catheter is curving and pointing towards the RV outflow tract, but sometimes in more dilated animals, this can change and the best loop can be when the catheter is directed towards the apex of the RV.

Getting negative pressures is not impossible. Depending on the situation, there can be some suction phenomenon inside the heart which can lead to negative pressures. Usually these are not very negative in a normal situation, and during disease minimum/diastolic pressures usually become higher.

We do not know, as this is the only protocol we have used. In our case, the choice of anesthesia was purely based on ease of access, and the fact that our anesthesiology team (clinical) was using it more frequently. I’m sure there are some papers comparing different anesthetics in PV data in rodents, and I think there might be also in pigs, but I can’t seem to find them again! If you know of such papers, please forward them to me!

So far, we’ve only used the 507 in pigs ranging from 40 to 90 kg. However, we have the feeling that we get better loops in smaller animals, which led us to get a 510 (10 mm electrode spacing) to test in the larger animals, which we didn’t get yet, so I can’t really comment on that. We can however get the 7 segments most of the times with the 507 in animals above 50 kg.

When we do it we place a short length introducer sheath and advance the Ventri Cath through it. This avoids any bleeding, and facilitates moving the catheter back and forth. However the hemostasis valve in a introducer sheath does not “fix” the catheter very well in place, so sometimes we need to provide some external support to keep it steady. Alternatively to that (and we have not done this, so I don’t know how easy it would be), you could introducer a short introducer into the LV, but this introducer would have to end in a “female luer” hub. To this hub you could adapt a Tuhoy borst valve, which would fix your catheter in place and make sure it would not move. Hope it helps!

During the acquisition always make sure you stop ventilation (when possible) and that you do not have arrhythmias. Most of the times, when you get arrhythmias during IVCO, it is because your catheter position is not ideal. If this is the case, then you are already getting bad data anyway. When my IVCO data has bad loops, I just have to accept it and not analyze it. That is why we always repeat IVCO several times, until we are sure we can get enough IVCO so that we can get some data.