Here we have presented a technique for measuring the axial contractile properties of immature heart cells, reporting the first purely axial force measurements of single developing cardiomyocytes. This general technique achieves single-cell manipulation in a parallel fashion with minimal labor and enables the study of both developing neonatal cardiomyocytes and stem cell-derived cardiomyocytes, which can be used in cardiac cell and tissue graft therapies. To create this system we modified a traditional force post assay (Tan et al. 2003
) by enlarging the posts and lattice constants and by filling the volume between the posts with a sacrificial layer of the biocompatible and thermoresponsive polymer, PNIPAAm. Previously PNIPAAm has been used as a release layer for creating contractile cell sheets (Okano et al. 1993
) and beating muscular actuators (Feinberg et al. 2007
) as well as a thick sacrificial layer for the creation of suspending multicellular cardiomyocyte constructs (Xi et al. 2005
). No patterning of protein or other adhesion material was required as with polyacrylamide gels (Théry et al. 2006
; Théry 2010
), standard force post arrays (Liu et al. 2010
), and multicellular cardiomyocyte systems (Xi et al. 2005
; Feinberg et al. 2007
). The aspect ratio (3:1) between large lattice constant and post diameter prescribed a physiological, elongated shape for the cardiomyocytes that has been shown to promote optimal sarcomeric alignment (Bray et al. 2008
). While optical measurement errors are problematic in studies with small micropost displacements, our system is less sensitive to the optical measurement errors.
As we have discussed, device calibration is critical to quantify and detect small changes in force generation across samples and populations. As seen in , large variation in micropost stiffness suggests that stiffness in microstructures may be more variable than bulk measurements. These micropost batch-to-batch stiffness variations may be caused by irregularities in bake temperature, time, and exposure to oxygen plasma. Post-to-post variations may be due to geometric irregularities in the microposts caused by mold degradation or imperfections in the original lithographic masks. Calibrations using piezoresistive cantilevers (Park et al. 2007
; Kim et al. 2011
) can be used to standardize cellular force transducer platforms and enable meaningful comparisons of measurements made using various techniques.
Absolute uncertainty in any given force measurement is the sum of the accuracy and precision. For studies in which the same cantilever was used for all calibrations, any offset errors due to accuracy are expected to be consistent. For these comparisons, the precision error due to post-to-post variation is the key parameter differentiating uncertainties between studies. When measurements using one technique are compared with measurements using other force sensors and calibrations, it is important to use the absolute uncertainty, because both the accuracy and precision will vary from system to system.
Previous experiments with adult cardiomyocytes and carbon fibers demonstrated peak isometric forces ranging from 2.42 μN (Yasuda et al. 2001
) to 5.7 μN (Nishimura et al. 2004
). Here we detected axial peak forces that were ten to one hundred times smaller. The axial forces generated by neonatal rat cardiomyocytes are smaller than the net forces measured on force post arrays with neonatal rat cardiomyocytes (Zhao and Zhang 2006
; Rodriguez et al. 2011
), but it is difficult to compare such measurements because in previous force post measurements the cells had spread out into shapes that are not physiologically realistic. These cardiomyocytes were applying forces in multiple axes and such irregularly spread shapes can exaggerate pre-stress and therefore enlarge contractile forces (Bollensdorff et al. 2011
). Without a simple axial loading scheme, forces cannot readily be compared.
Our measurements were carried out under auxotonic conditions (mimicking ejection of blood from the heart); as the cells contract against the micropost “springs,” they experience increasing load. Differences in loading condition may contribute to between-study discrepancies in the observed force magnitude. As reported previously,(Iribe et al. 2007
) isometric loading conditions (mimicking isovolumic contraction of the heart) in which the cell length is held constant during contraction produce larger forces than loaded contraction. If our devices were integrated into a micropost array stretching device (Bollensdorff et al. 2011
) to control cell length during contraction, we might observe larger forces. However, large forces have also been measured under auxotonic conditions; using novel polysilicon microgrippers, Lin et al. (Lin et al. 2000
) measured peak auxotonic loaded forces of 12.6 μN with device effective stiffness of 1.47 N/m, suggesting that loading condition alone is unlikely to underlie such a large discrepancy between immature and adult cardiomyocyte contractile forces. Therefore, the large difference between the magnitude of force produced by immature and adult heart cells may be due to biological differences between developing and adult cardiomyocytes.
There are several potential limitations in all of these studies. Cell shape, cell slippage, varying levels of initial pre-stress, and damage during handling may also contribute to this variation. By controlling cell shape, we expect that we have limited geometry-based variations in cell function. In addition, we did not observe cell slippage and believe slippage was not an issue for this study. Implementation of force feedback may enable initial pre-stress to be tuned and its effects studied, and standardization of the cell isolation and plating procedure may reduce differences due to subtle cell damage incurred during the isolation of immature cardiomyocytes and their subsequent plating onto the devices. Controlling for these factors should enable future studies to detect any biologically meaningful differences in contractile force generation between adult, neonatal, and stem cell-derived cardiomyocytes, and should highlight platform-dependent differences in cell health, adhesion, and slippage.
This approach provides cell-level functional information, so it is also an ideal complement for recent “microtissue” measurement platforms that utilize multicellular constructs of heart cells and extracellular matrix proteins (Feinberg et al. 2007
; Legant et al. 2009
; Grosberg et al. 2011
) to measure the active and passive properties of heart-like tissue. Both platforms could be used in conjunction to study the role of cell-level mutations and tissue composition and organization in disease progression.
Future work should address sources of the differences in the forces generated by immature and adult cardiomyocytes. It will be important to identify potential cellular differences in behavior for this assay compared to other two-point methods. Additionally, future studies should address changes in contractile force generation as a function of cell age, morphology, sarcomere organization, and external environmental parameters. Ultimately we would like to be able to perform studies of immature and adult cardiomyocytes with our platform. Adhesion is extremely challenging with adult cardiomyocytes. The few micropost studies that have achieved it have required at least 24 hours in culture, and these studies have observed significant dedifferentiation and unhealthy contractility changes (Zhao and Zhang 2006
; Zhao et al. 2007
). For future studies we aim to develop a protocol for robustly adhering adult heart cells to microposts in less than an hour and thereby enable contractile measurements of “healthy” adult cardiomyocytes. We have reported the first steps towards this goal using a PNIPAAm-free adhesion process (Taylor et al. 2011
This technique may also be applicable to other tension-sensitive cell types (such as neurons and fibroblasts), and to investigate changes in force production over time in healthy and diseased cells. Finally, this sacrificial layer technique is broadly applicable as a highly parallel technique for controlling cell placement and shape. We anticipate that our method will be suitable for tissue engineering applications such as co-culture on 3D topographies as well as high-throughput pharmaceutical screening.