WO2016191179A1 - In vitro models of regenerative muscle cell therapy and uses thereof - Google Patents

Abstract

The present invention provides in vitro models of regenerative muscle cell therapy and uses thereof to determine the ability of a stem cell to functionally engraft with a myocyte, e.g., the functional coupling of native and stem cell-derived myocytes.

Description

IN VITRO MODELS OF REGENERATIVE MUSCLE CELL THERAPY AND USES

THEREOF

RELATED APPLICATIONS

This application claims priority to U.S. Provisional Patent Application No.62/165,371, filed on May 22, 2015, the entire contents of which are incorporated herein by this reference.

GOVERNMENT SUPPORT

This invention was made with government support under grant number UH3 TR000522, awarded by NIH/NCATS and grant number U01 HL100408 awarded by NIH/NHLBI. The government has certain rights in the invention.

BACKGROUND OF THE INVENTION

The goal of stem cell transplantation therapy is to improve clinical outcomes in the treatment of, e.g., muscular disease. For example, cardiac stem cell therapy aims at regenerating lost muscle tissue in the heart by administering to a subject various types of stem and progenitor cells. Despite a positive safety outcome (Menasche P, et al. (2001) Lancet 357(9252):279-280; Strauer BE, et al. (2002) Circulation 106(15):1913-1918; Makkar RR, et al. (2012) Lancet 379(9819):895-904), however, stem cell transplantation clinical trials, e.g., cardiac stem cell transplantation clinical trials, have shown inconsistent improvements in cardiac function, ranging from no recovery at all (Makkar RR, et al. (2012) Lancet 379(9819):895-904; Marban E & Malliaras K (2012) JAMA 308(22):2405-2406;Traverse JH, et al. (2012) JAMA 308(22):2380- 2389) to between 5% and 10% increase in left ventricular ejection fraction (Abdel-Latif A, et al. (2007) Archives Int Med 167(10):989-997; Donndorf P, et al. (2011) J Thorac Cardiovasc Surg 142(4):911-920; Bolli R, et al. (2011) Lancet 378(9806):1847-1857). Efficacy was limited by poor stem cell engraftment and survival, with cardiac retention rates of less than 1% at twenty- four hours post transplantation and stem cell accumulation in other organs (Dow J, et al. (2005) Cardiovascular research 67(2):301-307; Penicka M, et al. (2005) Circulation 112(4):e63-65; Muller-Ehmsen J, et al. (2006) J Mol Cell Cardiol 41(5):876-884; Hudson W, et al. (2007) J Surg Res 142(2):263-267). At the same time, preliminary studies of embryonic stem cells applied to whole heart preparations indicated potential for stem cell engraftment to the myocardium, with some degree of electrical coupling (Kehat I, et al. (2004) Nat Biotech 22(10):1282-1289.; Shiba Y, et al. (2012) Nature 489(7415):322-325), but only partial augmentation of contractile function was reported (Hudson W, et al. (2007) J Surg Res 142(2):263-267; Kolossov E, et al. (2006) J Exp Med 203(10):2315-2327; Laflamme MA, et al. (2007) Ann Rev Pathol 2:307-339.).

The myocardium is a functional syncytium of seamlessly integrated myocytes. For transplanted stem cells to effectively couple with the native tissue, they must organize myofibrils and form junctions to adjacent cells, known as intercalated disks, through which electro-chemical signals and mechanical forces are transmitted. The co-ordinated assembly of these structures relies on the distribution and remodeling of cell-matrix and cell-cell adhesions between stem cells and existing myocytes (Woods JR, Jr., et al. (1978) Am J Physiol 234(5):H520-524; Hirschy A, et al. (2006) Dev Biol 289(2):430-441; Wu JC, et al. (2002) J Cell Biochem 84(4):717-724;

McCain ML, et al. (2012) PNAS, USA 109(25):9881-9886) that guide the alignment of myofibrils to establish the directionality of cardiac contraction (Parker KK & Ingber DE (2007) Philo Trans Roy Soc London Series B, Biological sciences 362(1484):1267-1279; Feinberg AW, et al. (2012) Biomaterials 33(23):5732-5741; Feinberg AW, et al. (2007) Science 317(5843):1366-1370; Grosberg A, et al. (2011) PLoS Comp Biol 7(2):e1001088.). Furthermore, intercalated discs contain gap junctions that mediate electro-chemical coupling and orchestrate synchronous force generation (Jansen JA, et al. (2010) J Mol Cell Cardiol 48(1):76-82; Bers DM (2002) Nature 415(6868):198-205). Thus, for stem cell therapy to be effective, newly formed cells must not only be structurally similar to native myocytes, but must also form mechanical and electrical cell-cell junctions with the native tissue to ensure complete electrical and mechanical synchrony.

Accordingly, there is a need in the art for methods to identify suitable cells for myocyte regeneration therapy, such as myocyte stem cell transplantation therapy, agents that are useful for tissue engraftment of a stem cell to native tissue, agents useful for regenerative muscle cell therapy, and devices for performing the same.

SUMMARY OF THE INVENTION

The present invention is based, at least in part, on the discovery of devices and methods for determining the ability of a stem cell to functionally engraft, e.g., mechanically and electrically, couple, with a myocyte, such as a cardiomyocyte, in order to, e.g., identify cells and agents suitable for use in regenerative muscle cell therapy. The devices and methods described herein permit an in vitro determination of the ability of a stem cell to functionally engraft with a myocyte on a spatial and temporal resolution scale that is superior to existing in vivo approaches. In particular, devices which recapitulate the in vivo milieu of myocytes, e.g., soft substrates comprising a patterned biopolymer, were developed, and using murine post-natal cardiomyocytes, e.g., neonate ventricular myocytes (e.g.,“mature cells”), and immature embryonic (mES-) or induced pluripotent stem cell-derived myocytes (miPS-CMs), it has been discovered that the cells structurally integrate as revealed by the presence of aligned actin myofibrils and β-catenin and connexin 43 containing cell junctions between neonate and stem cell-derived myocytes. Ratiometric Ca2+ imaging and traction force microscopy revealed synchronous Ca2+ transients and mechanical contractions between cells, but reduced Ca2+ levels and lower peak systolic forces were observed in mES- and miPS-CMs coupled with neonate myocytes. These differences yield an imbalance in tension across the cells that was accompanied by the appearance of traction forces and substrate adhesions near the cell-cell junction. A finite element model of muscle contraction revealed that differences in isometric tension were sufficient to predict the observed pattern of adhesive forces on the substrate. Thus, despite achieving synchronous contraction, reduced force transmission between spared and newly formed myocytes limits repair of the contractile function in cardiac cell therapy.

Accordingly, in one aspect, the present invention provides in vitro methods for determining the ability of a stem cell to functionally engraft with a myocyte. The methods include providing a cell co-culture, comprising a base layer; a soft substrate on the base layer; a biopolymer patterned on the soft substrate; a single myocyte; and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface; determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, and identifying the stem cell as suitable for functionally engrafting with the myocyte if the contractile force of the stem cell is about the same as the contractile force of the myocyte , thereby determining the ability of the stem cell to functionally engraft with the myocyte .

In another aspect, the present invention provides in vitro methods for identifying a compound useful for functionally engrafting a stem cell and a myocyte. The methods include providing a cell co-culture, comprising a base layer; a soft substrate on the base layer; a biopolymer patterned on the soft substrate; a single myocyte; and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface; contacting the co-culture with a test compound; and determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for functionally engrafting the myocyte and the stem cell, thereby identifying a compound useful for functionally engrafting the stem cell and the myocyte.

In another aspect, the present invention provides in vitro methods for identifying a compound useful for treating a subject having regenerative muscle cell therapy. The methods include providing a cell co-culture, comprising a base layer; a soft substrate on the base layer; a biopolymer patterned on the soft substrate; a single myocyte; and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface; contacting the co-culture with a test compound; and determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for treating a subject having regenerative muscle cell therapy, thereby identifying a compound useful for treating a subject having regenerative muscle cell therapy.

The stem cell may be selected from the group consisting of a primary embryonic stem cell, a stem cell from an embryonic stem cell line, a primary fetal stem cell, a stem cell from a fetal stem cell line, a primary adult stem cell, a stem cell from an adult stem cell line, a stem cell de-differentiated from an adult cell, and an induced pluripotent stem cell (iPS).

The myocyte may be selected from the group consisting of a cardiomyocyte, a vascular smooth muscle cell, a smooth muscle cell, a skeletal striated muscle cell, and an airway smooth muscle cell.

The soft substrate may have a Young’s modulus of about 1 to about 100,000 pascal (Pa); or a Young’s modulus of about 1 to about 25,000 pascal (Pa).

The soft substrate may be selected from the group consisting of a polyacrylamide gel, a poly(N-isopropylacrylamide), a pHEMA, a collagen, a fibrin, a gelatin, an alginate, and a dextran.

The base layer may be selected from the group consisting of a silicon wafer, a glass cover slip, a multi-well plate, a tissue culture plate, and a chamber of a microfluidic device.

The biopolymer may be selected from the group consisting of a protein, a carbohydrate, a lipid, and a nucleic acid, or combinations thereof. In one embodiment, the biopolymer is selected from the group consisting of a fibronectin, a laminin, a collagen, a gelatin, a fibrin, a vitronectin, and a fibrinogen, and combinations thereof. In another embodiment, the biopolymer is fibronectin.

The soft substrate may functionalized with a bifunctional or a multifunctional linker molecule. In one embodiment, the bifunctional or multifunctional linker is N-sulfosuccinimidyl 6-hexanoate (sulfo-SANPAH). In another embodiment, the soft substrate is functionalized with a repellent, e.g.,agarose, hyaluronic acid, or alginate.

The soft substrate may further comprise a fluorescent bead, such as a fluorescent bead embedded within the soft substrate.

In one aspect, the present invention provides in vitro model of regenerative muscle cell therapy. The model includes a cell co-culture, comprising a base layer; a soft substrate on the base layer; a biopolymer patterned on the soft substrate; a single myocyte ; and a single stem cell, wherein the single myocyte and the single stem cell are cultured such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface.

BRIEF DESCRIPTION OF THE DRAWINGS

Figure 1Ai shows representative images of mono-nucleated (DAPI) mouse neonate, mES- or miPS- cardiac myocytes (CMs) stained for actin. Scale 10 µm.

Figure 1Aii shows representative images of mono-nucleated (DAPI) mouse neonate, mES- or miPS- cardiac myocytes (CMs) stained for actin. Scale 10 µm.

Figure 1B shows actin orientation order parameter (OOP) as a function of cell type (n = 25, 20 and 9 for Neonate, mES, and miPS, respectively).

Figure 1Ci shows representative heat maps indicating peak systolic displacement.

Figure 1Cii shows representative heat maps indicating traction stress.

Figure 1D displays peak systolic force as a function of cell type (n = 8, 7 and 7 for Neonate, mES, and miPS, respectively). Results presented as mean ±sem. * indicates statistically significant difference with a p-value <0.05.

Figure 2A shows representative images of pairs of mono-nucleated (DAPI) mouse neonate, mES or miPS-CMs stained for actin. Scale 10 µm. GFP-tagged mES or miPS-CMs were used for heterogeneous pairing.

Figure 2B shows representative images of pairs of mono-nucleated (DAPI) mouse neonate, mES or miPS-CMs stained for β-catenin. Scale 10 µm. GFP-tagged mES or miPS-CMs were used for heterogeneous pairing. Figure 2C shows representative images of pairs of mono-nucleated (DAPI) mouse neonate, mES or miPS-CMs stained for connexin-43. Scale 10 µm. GFP-tagged mES or miPS- CMs were used for heterogeneous pairing.

Figure 2D shows representative ratiometric Ca²⁺ transients across several contraction cycles for different µtissue types.

Figure 2Ei is a graph showing the normalized cross-correlation functions for different µtissue types.

Figure 2Eii is a graph showing the normalized cross-correlation coefficients for different µtissue types.

Figure 2F shows diastolic Ca²⁺ levels for each cell within a cell pair. Results are presented as mean ±sem (n = 8, 13, 9, 15 and 6 for Neonate-Neonate, mES-mES, mES-Neonate, miPS-miPS, miPS-Neonate, respectively). * indicates statistically significant difference with homogeneous neonate pair (p <0.05).

Figure 3A shows representative heat maps indicating peak systolic displacement (Figure 3Ai, traction stress (Figure 3Aii) for pairs of mono-nucleated (DAPI) myocytes. Scale 10µm.

Figure 3B shows representative vinculin-positive adhesions (arrows) for pairs of mono- nucleated (DAPI) myocytes. Scale 10µm.

Figure 3Ci shows schematic diagram indicating force calculations for different locations within cell pairs.

Figure 3Cii shows representative contraction cycles for each cell within homogeneous cell pairs.

Figure 3Ciii shows representative contraction cycles for each cell within heterogeneous mES-CMs cell pairs.

Figure 3Civ shows representative contraction cycles for each cell within

heterogeneousmiPS-CMs (Figure 3Civ) cell pairs.

Figure 3D shows the duration of the contraction cycle as a function of micro-tissue type. Figure 3E displays peak systolic force in different regions within cell pairs.

Figure 3F shows the percent cell shortening across homogeneous and heterogeneous cell pairs. Results presented as mean ±sem (n = 7, 12, 8, 7 and 7 for Neonate-Neonate, mES-mES, mES-Neonate, miPS-miPS and miPS-Neonate, respectively). Statistically significant differences (p<0.05) from the neonate-neonate case are indicated with * for cellular values and† for junctional ones.

Figure 4A shows representative finite element mesh of micro-tissue (cell-cell junction). Figure 4B shows the predicted patterns of local traction for (Figure 4Bi) homogeneous and (Figure 4Bii) heterogeneous pairs.

Figure 4Ci shows predicted patterns of cellular stress at peak systole for (Figure 4Ci) homogeneous cell pairs. The gray arrow indicates high traction stress beneath the neonate myocyte near the cell-cell junction.

Figure 4Cii shows predicted patterns of cellular stress at peak systole for (Figure 4Ci) heterogeneous cell pairs. The gray arrow indicates high traction stress beneath the neonate myocyte near the cell-cell junction.

Figure 4D shows predicted traction force and percent shortening in heterogeneous cell pairs.

Figure 4Ei is a schematic illustrating the typical tension experienced at substrate adhesions within pairs of neonate cells µtissues.

Figure 4Eii is a schematic illustrating the typical tension experienced at substrate adhesions within pairs of heterogeneousµtissues.

Figure 5A shows the surface area for Neonate-Neonate, mES-mES, mES-Neonate, miPS- miPS and miPS-Neonate (n = 14, 9, 12, 8 and 9, respectively).

Figure 5B shows Actin Orientation Order Parameter for Neonate-Neonate, mES-mES, mES-Neonate, miPS-miPS and miPS-Neonate (n = 8, 8, 6, 4 and 8, respectively).

Figure 5C displays a slope of the cell-cell junction for Neonate-Neonate, mES-mES, mES-Neonate, miPS-miPS and miPS-Neonate n = 11, 6, 10, 6 and 9, respectively).

Figure 5D displays the nuclear eccentricity (n = 5 for all µtissues). Results presented as mean ±sem, statistical differences (p<0.05) is indicated with *.

Figure 6A shows DIC (top) and traction force field (bottom) images of mES-derived and neonate cardiomyocytes seeded on fibronectin-coated islands with length to width ratios of 3:1, mimicking concentric and eccentric hypertrophy, respectively.

Figure 6B shows DIC (top) and traction force field (bottom) images of mES-derived and neonate cardiomyocytes seeded on fibronectin-coated islands with length to width ratios of 9:1 (Figure 6B), mimicking concentric and eccentric hypertrophy, respectively.

Figure 6C shows the DIC (left) and traction force field (right) images of mES-derived and neonate cardiomyocytes seeded on fibronectin islands with length to width ratios of 7:1 micro- contact printed on a stiff (90 kPa) substrate, representing a fibrotic microenvironment. Figure 6D displays summary statistics that show the percentage of the peak systolic traction force dissipated at the cell-cell junction in pathological (3:1, 9:1, 7:1 at 90kPa) conditions is similar or greater than observed in physiological (7:1 at 13kPa) conditions (gray box, main Figure 2). Results are mean ±sem (n = 13, 6 and 7 for 3:1, 9:1 and 90 kPa, respectively). * indicates statistically significant difference with p <0.05.

Figure 7A shows a section of the micro-tissue depicting the cellular tension along the longitudinal direction and the traction stress on the gel substrate.

Figure 7B displays a Hill’s three-element model for the muscle contraction.

Figure 7C shows the linearized stress-strain rate relation (i.e., the inset equation) for the contractile element of Hill’s three-element model.

Figure 7D shows a schematic for determining the cell-substrate linkage constant at the steady state using a positive feedback between the focal adhesion maturation and traction stress.

DETAILED DESCRIPTION OF THE INVENTION

The present invention is based, at least in part, on the discovery of devices and methods for determining the ability of a stem cell to functionally engraft, e.g., mechanically and electrically, couple, with a myocyte, such as a cardiomyocyte, in order to, e.g., identify cells and agents suitable for use in regenerative muscle cell therapy. The devices and methods described herein permit an in vitro determination of the ability of a stem cell to functionally engraft with a myocyte on a spatial and temporal resolution scale that is superior to existing in vivo approaches. In particular, devices which recapitulate the in vivo milieu of myocytes, e.g., soft substrates comprising a patterned biopolymer, were developed, and using murine post-natal

cardiomyocytes, e.g., neonate ventricular myocytes (e.g.,“mature cells”), and immature embryonic (mES-) or induced pluripotent stem cell-derived myocytes (miPS-CMs), it has been discovered that the cells structurally integrate as revealed by the presence of aligned actin myofibrils and β-catenin and connexin 43 containing cell junctions between neonate and stem cell-derived myocytes. Ratiometric Ca2+ imaging and traction force microscopy revealed synchronous Ca2+ transients and mechanical contractions between cells, but reduced Ca2+ levels and lower peak systolic forces were observed in mES- and miPS-CMs coupled with neonate myocytes. These differences yield an imbalance in tension across the cells that was accompanied by the appearance of traction forces and substrate adhesions near the cell-cell junction. A finite element model of muscle contraction revealed that differences in isometric tension were sufficient to predict the observed pattern of adhesive forces on the substrate. Thus, despite achieving synchronous contraction, reduced force transmission between spared and newly formed myocytes limits repair of the contractile function in cardiac cell therapy.

I. Methods of the Invention

Provided herein are in vitro methods for determining the ability of a stem cell to functionally engraft with a myocyte, e.g., a cardiomyocyte. Also provided herein are screening methods to identify compounds useful for functionally engrafting a myocyte, e.g., a

cardiomyocyte, and a stem cell, and methods for identifying compounds useful for treating a subject having regenerative muscle cell therapy. The methods of the invention generally include providing a co-culture comprising a single myocyte, e.g., a mature myocyte, and a single stem cell and determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell.

In one aspect, the present invention provides methods for determining the ability of a stem cell to functionally engraft with a myocyte. The methods include providing a cell co-culture comprising a base layer, a soft substrate on the base layer, a biopolymer patterned on the soft substrate, a single myocyte, e.g., a mature myocyte, and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell-to- cell interface; determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell; identifying the stem cell as suitable for functionally engrafting with a myocyte if the contractile force of the stem cell is about the same as the contractile force of the myocyte , thereby determining the ability of the stem cell to functionally engraft with the myocyte .

The present invention also provides in vitro methods for identifying a compound useful for functionally engrafting a stem cell and a myocyte. The methods include providing a cell co- culture comprising a base layer, a soft substrate on the base layer, a biopolymer patterned on the soft substrate, a single myocyte, e.g., a mature myocyte, and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell- to-cell interface; contacting the co-culture with a test compound; and determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell in the presence of the test compound and in the absence of the test compound, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for functionally engrafting a stem cell and a myocyte, thereby identifying a compound useful for functionally engrafting a stem cell and a myocyte. The present invention further provides methods for identifying a compound useful for treating a subject having regenerative muscle cell therapy. The methods include providing a cell co-culture comprising a base layer, a soft substrate on the base layer, a biopolymer patterned on the soft substrate, a single myocyte, e.g., a mature myocyte, and a single stem cell; culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell- to-cell interface; contacting the co-culture with a test compound; and determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell in the presence of the test compound and in the absence of the test compound, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for treating a subject having regenerative muscle cell therapy, thereby identifying a compound useful for treating a subject having regenerative muscle cell therapy.

In further aspects, the methods of the present invention can be used to study functional differentiation of stem cells (e.g., pluripotent stem cells, multipotent stem cells, induced pluripotent stem cells, and progenitor cells of embryonic, fetal, neonatal, juvenile and adult origin) into contractile phenotypes or dedifferentiation of mature cells, e.g., differentiated cells, into less mature cell types.

The articles“a” and“an” are used herein to refer to one or to more than one (i.e., to at least one) of the grammatical object of the article. By way of example,“an element” means one element or more than one element, e.g., a plurality of elements.

The term“including” is used herein to mean, and is used interchangeably with, the phrase “including but not limited to”.

The term“or” is used herein to mean, and is used interchangeably with, the term “and/or,” unless context clearly indicates otherwise.

The term“at least” prior to a number or series of numbers is understood to include the number adjacent to the term“at least”, and all subsequent numbers or integers that could logically be included, as clear from context. When at least is present before a series of numbers or a range, it is understood that“at least” can modify each of the numbers in the series or range.

As used herein, ranges include both the upper and lower limit. The term“about” is used herein to mean within the typical ranges of tolerances in the art. For example,“about” can be understood as about 2 standard deviations from the mean. When about is present before a series of numbers or a range, it is understood that“about” can modify each of the numbers in the series or range. When referring to a measurable value such as an amount, a temporal duration, and the like, the term“about” is meant to encompass variations of ±20%, ±10%, ±5%, ±1%, and ±0.1% from the specified value, as such variations are appropriate.

The term“engraft” as used herein refers to coupling of two cells from different sources, such as a stem cell and a myocyte. The cells, e.g., a stem cell and a myocyte, may structurally engraft, functionally engraft, or structurally and functionally engraft.

As used herein, the term“structurally engraft,” used interchangeable with the term “structurally integrate,” when referring to a stem cell and a myocyte, indicates that the myofibrils of the cells have aligned parallel to the long axis of the cell, and/or that the myofibrils cross cell- to-cell junctions, and/or that the cells maintain a registered array or sarcomeres, and/or that the cells form cell-to-cell gap junctions and/or cell-to-cell adherens junctions.

Methods to determine if a stem cell and a myocyte have structurally integrated, include microscopic analyses, including but not limited to fluorescent microscopy, confocal microscopy, two-photon microscopy, and the like, immunohistochemical analyses, e.g., staining for connexin 43 to determine if the cells have formed electrically-competent junctions, staining for β-catenin to determine if the cells have formed mechanically-competent junctions, staining for α-actin and determining, e.g., the orientational order parameter (OOP) of the networks to determine if the cells have formed registered myofibrils.

As used herein, the term“functionally engraft,” used interchangeable with the term “functionally integrate,” when referring to a stem cell and a myocyte, indicates that the cells have mechanically and electrically integrated, e.g., the cells synchronously contract, and/or the cells generate a contractile force, and/or the contractions of the cells are in phase, and/or the contractile force at the medial cell-to-cell junction of the stem cell and the myocyte is about the same. A stem cell and a myocyte can further be identified as having functionally engrafted if, e.g., the cells exhibit synchronous Ca⁺² transients, and/or the cells exhibit substantially the same Ca⁺² levels, and/or the cells exhibit peak systolic and/or diastolic forces that are about the same.

Methods to determine if a stem cell and a myocyte have functionally integrated, include microscopic analyses, including but not limited to fluorescent microscopy, confocal microscopy, two-photon microscopy, and the like, immunohistochemical analyses, e.g., vinculin staining, traction force microscopy, ratiometric Ca⁺² imaging, and optical mapping of the action potentials. In one embodiment, the methods of the invention include determining the contractile force of the stem cell and the myocyte at the medial cell-to-cell interface using traction force microscopy. In such embodiments, the soft substrate may further include a fluorescent bead and the displacement of the fluorescent bead is determined.

The term“stem cell” as used herein, refers to an undifferentiated cell which is capable of proliferation and giving rise to more progenitor cells having the ability to generate a large number of mother cells that can in turn give rise to differentiated, or differentiable daughter cells. The daughter cells themselves can be induced to proliferate and produce progeny that subsequently differentiate into one or more mature cell types, while also retaining one or more cells with parental developmental potential. The term“stem cell” refers to a subset of progenitors that have the capacity or potential, under particular circumstances, to differentiate to a more specialized or differentiated phenotype, and which retains the capacity, under certain circumstances, to proliferate without substantially differentiating. In one embodiment, the term stem cell refers generally to a naturally occurring mother cell whose descendants (progeny) specialize, often in different directions, by differentiation, e.g., by acquiring completely individual characters, as occurs in progressive diversification of embryonic cells and tissues. Cellular differentiation is a complex process typically occurring through many cell divisions. A differentiated cell may derive from a multipotent cell which itself is derived from a multipotent cell, and so on. While each of these multipotent cells may be considered stem cells, the range of cell types each can give rise to may vary considerably. Some differentiated cells also have the capacity to give rise to cells of greater developmental potential. Such capacity may be natural or may be induced artificially upon treatment with various factors. In many biological instances, stem cells are also

“multipotent” because they can produce progeny of more than one distinct cell type, but this is not required for“stem-ness.” Self-renewal is the other classical part of the stem cell definition. In theory, self-renewal can occur by either of two major mechanisms. Stem cells may divide asymmetrically, with one daughter retaining the stem state and the other daughter expressing some distinct other specific function and phenotype. Alternatively, some of the stem cells in a population can divide symmetrically into two stems, thus maintaining some stem cells in the population as a whole, while other cells in the population give rise to differentiated progeny only. Formally, it is possible that cells that begin as stem cells might proceed toward a differentiated phenotype, but then“reverse” and re-express the stem cell phenotype, a term often referred to as “dedifferentiation” or“reprogramming” or“retrodifferentiation”. The term“embryonic stem cell” is used to refer to the pluripotent stem cells of the inner cell mass of the embryonic blastocyst (see, e.g., U.S. Patent Nos.5,843,780, 6,200,806, the entire contents of each of which are incorporated herein by reference). Such cells can similarly be obtained from the inner cell mass of blastocysts derived from somatic cell nuclear transfer (see, for example, U.S. Patent Nos.5,945,577, 5,994,619, 6,235,970, the entire contents of each of which are incorporated herein by reference). The distinguishing characteristics of an embryonic stem cell define an embryonic stem cell phenotype. Accordingly, a cell has the phenotype of an embryonic stem cell if it possesses one or more of the unique characteristics of an embryonic stem cell such that that cell can be distinguished from other cells. Exemplary distinguishing embryonic stem cell characteristics include, without limitation, gene expression profile, proliferative capacity, differentiation capacity, karyotype, responsiveness to particular culture conditions, and the like.

The term“adult stem cell” or“ASC” is used to refer to any multipotent stem cell derived from non-embryonic tissue, including fetal, juvenile, and adult tissue. Stem cells have been isolated from a wide variety of adult tissues including blood, bone marrow, brain, olfactory epithelium, skin, pancreas, skeletal muscle, and cardiac muscle. Each of these stem cells can be characterized based on gene expression, factor responsiveness, and morphology in culture.

Exemplary adult stem cells include neural stem cells, neural crest stem cells, mesenchymal stem cells, hematopoietic stem cells, and pancreatic stem cells.

The term“progenitor cell” is used herein to refer to cells that have a cellular phenotype that is more primitive (e.g., is at an earlier step along a developmental pathway or progression than is a fully differentiated cell) relative to a cell which it can give rise to by differentiation. Often, progenitor cells also have significant or very high proliferative potential. Progenitor cells can give rise to multiple distinct differentiated cell types or to a single differentiated cell type, depending on the developmental pathway and on the environment in which the cells develop and differentiate. Furthermore, the term“progenitor cell” is used herein synonymously with“stem cell.”

In one embodiment, progenitor cells suitable for use in the claimed devices and methods are Committed Ventricular Progenitor (CVP) cells as described in PCT Application No. WO 2010/042856, entitled“Tissue Engineered Mycocardium and Methods of Productions and Uses Thereof,” filed October 9, 2009, the entire contents of which are incorporated herein by reference. Suitable stem cells for use in the present invention are stem cells that will differentiate into a myocyte, the differentiated progeny of such stem cells, and the dedifferentiated progeny of myocytes, and include embryonic (primary and cell lines), fetal (primary and cell lines), adult (primary and cell lines) and iPS (induced pluripotent stem cells). The stem cells may be normal stem cells, abnormal stem cells (e.g., those derived from a diseased tissue, or those that are physically or genetically altered to achieve an abnormal or pathological phenotype or function), normal or diseased cells derived from embryonic stem cells or induced pluripotent stem cells, or cells comprising a genetic construct, such as an expression vector expressing an optogenetic gene, e.g., a light sensitive ion channel (e.g., channelrhodopsin (ChR2), C1V1, Chrimson, Chronos, SSFO, ArchT, ChETA, NpHR, SwiChR, iC1C2, or the like).

Stem cells can be cultured in vitro, derived from a natural source, genetically engineered, or produced by any other means. Any natural source of cells may be used. For example, in one embodiment the stem cells suitable for use in the present invention, e.g., stem cells that give rise to myocytes, may be selected from the group consisting of a primary embryonic stem cell, a stem cell from an embryonic stem cell line, a primary fetal stem cell, a stem cell from a fetal stem cell line, a primary adult stem cell, a stem cell from an adult stem cell line, a stem cell de- differentiated from an adult cell, and an induced pluripotent stem cell (iPS).

As used herein, the term“myocyte” or“mature myocyte” refers to a cell exhibiting structural and functional signatures of post-natal myocytes. For example, in the case of cardiomyocytes, these signatures may be structural and include, for example, the presence of a contractile cytoskeleton characterized by well-oriented, and well-organized sarcomeres and/or greater sarcomere length and higher sarcomere packing density of myocytes as compared to mES or miPS stem cells, and/or an orientational order parameter (OOP) of myofibril networks about higher than an OOP of mES or miPS stem cells. Methods and reference standards for the structural signatures of myocytes are described in U.S. Provisional Patent Application No.

62/118,806 filed on February 20, 2015, the entire contents of which are incorporated herein by reference. Furthermore, functional signals also typify myocytes, e.g., cardiomyocytes. For example, a negative resting membrane potential, a fast and robust action potential upstroke (e.g., stem cells, such as mES and miPS stem cells exhibit equal proportion if ventricular-like and atrial-like action potential traces while myocytes exhibit primarily ventricular-like action potential traces as determined by patch clamp recording), appropriate duration at 30, 50, 70, and 90% of the action potentials calcium transient, and the twitch stress and relaxation time for the contractile cycle (e.g., Ca2+ transients measurements have a significantly (p<0.05) shorter 50% decay time (CaT50) in the miPS, but not the mES tissues, as compared to the neonate, and significantly (p<0.05) shorter 90% decay time (CaT90) in both the mES and miPS cells versus the neonate tissues; L- and T- type Ca2+ current profiles are significantly (p<0.05) higher total (TOT) and T-type (TCC) maximum Ca2+ current densities in the neonate myocytes versus the mES-derived, but not the miPS-derived myocytes). Methods and reference standards for the functional signatures of myocytes are also described in U.S. Provisional Patent Application No. 62/118,806 filed on February 20, 2015.

The myocytes may be normal myocytes, abnormal myocytes (e.g., those derived from a diseased tissue, or those that are physically or genetically altered to achieve an abnormal or pathological phenotype or function), normal or diseased myocytes derived from embryonic stem cells or induced pluripotent stem cells, or myocytes comprising a genetic construct, such as an expression vector expressing an optogenetic gene, e.g., a light sensitive ion channel (e.g., channelrhodopsin (ChR2), C1V1, Chrimson, Chronos, SSFO, ArchT, ChETA, NpHR, SwiChR, iC1C2, or the like).. Myocytes can be cultured in vitro, derived from a natural source, genetically engineered, or produced by any other means. Any natural source of myocytes may be used, including from neonates, e.g., mouse and rat neonates.

The myocytes for use in the present invention may be cardiomyocytes, vascular smooth muscle cells, smooth muscle cells, skeletal striated muscle cells, or airway smooth muscle cells.

As used herein, the term“co-culture” refers to a combined cell culture of more than one distinct cell type.

As used herein, the various forms of the term "modulate" are intended to include stimulation (e.g., increasing or upregulating a particular response or activity) and inhibition (e.g., decreasing or downregulating a particular response or activity).

As used herein, the term "contacting" (e.g., contacting cells seeded on a soft substrate with a test compound) is intended to include any form of interaction (e.g., direct or indirect interaction) of a test compound and cells on or within a soft substrate. The term contacting includes incubating a compound and cells on or within a soft substrate (e.g., adding the test compound to cells).

In one embodiment, the methods of the invention further include applying a stimulus, such as an electrical stimulus or a chemical stimulus, or, in the case of cells expressing an optogenetic gene, a light stimulus, to the cells.

Test compounds, may be any agents including chemical agents (such as toxins), small molecules, pharmaceuticals, peptides, proteins (such as antibodies, cytokines, enzymes, and the like), and nucleic acids, including gene medicines and introduced genes, which may encode therapeutic agents, such as proteins, antisense agents (i.e., nucleic acids comprising a sequence complementary to a target RNA expressed in a target cell type, such as RNAi or siRNA), ribozymes, and the like.

The test compound may be added to cells by any suitable means. For example, the test compound may be added drop-wise onto the surface of the cells and allowed to diffuse into or otherwise enter the cells, or it can be added to the nutrient medium and allowed to diffuse through the medium. In the embodiment where the cell cells on a soft substrate is cultured in a multi-well plate, each of the culture wells may be contacted with a different test compound or the same test compound. In one embodiment, the screening platform includes a microfluidics handling system to deliver a test compound and simulate exposure of the microvasculature to drug delivery.

In certain embodiments, the methods of the invention are high throughput methods, where a plurality of test compositions or conditions are screened. For example, in certain embodiments, a library of compositions are screened, where each composition of the library is individually contacted to the co-cultures in order to identify which agents suitable for use as described herein.

Subjects having regenerative muscle cell therapy include, for example, subjects having an acute myocardial infarction, subjects recovering from a recent myocardial infarction, subjects with pre-existing ischemic damage that is leading to heart failure, or subjects suffering from heart failure in the early or late phase of the disease. Ischemic diseases include, for example, ischemic disease of cardiac muscle and limb ischemia. Additional examples of subjects having

regenerative muscle cell therapy include, for example, subjects having a muscular dystrophy, such as Duchenne Muscular Dystrophy, and subjects having the morbidities and co-morbidities associated with diabetes.

The methods of the present invention may further comprise, e.g., imaging the cells on the soft substrate and/or staining the cells on the soft substrate for a particular cell type or gene or protein expression. Imaged cells can be analyzed using structural phenotyping methods and metrics designed to assess the quality of stem cell-derived myocytes described in, for example, U.S. Provisional Patent Application No.62/118,806, filed on February 20, 2015.

The methods of the present invention may also further include determining an electrophysiological activity of the cells, such as, for example, a voltage parameter selected, e.g., an action potential, action potential duration (APD), conduction velocity (CV), refractory period, wavelength, restitution, and/or a calcium flux parameter, e.g., intracellular calcium transient, transient amplitude, rise time (contraction), decay time (relaxation), total area under the transient (force), restitution, focal and spontaneous calcium release. The methods of the invention may further include determining the health and maturation of mitochondria and other intracellular organelles using, e.g., fluorescent dyes and live microscopy.

Gene expression can be also further be determined in the methods of the invention using, e.g., FISH and other multiplexed techniques such as FISSEQ.

In certain embodiments of the invention, e.g., for evaluation of electrophysiological activities, cells may be co-cultured in the presence of a fluorophor such as a voltage-sensitive dye or an ion-sensitive dye. For example, the voltage-sensitive dye is an electrochromic dye such as a a styryl dye or a merocyanine dye. Exemplary electrochromic dyes include RH-421 or di-4- ANEPPS. Ion-sensitive, e.g., calcium sensitive dyes, include aequorin, Fluo3, and Rhod2. For simultaneous measurements of action potentials and intracellular calcium, the following exemplary dye pairs are used: di-2-ANEPEQ and calcium green; di-4-ANEPPS and Indo 1; di-4- ANEPPS and Fluo-4; RH237 and Rhod2; and, RH-237 and Fluo-3/4.

The methods and devices of the invention are also useful for evaluating the effects of particular delivery vehicles for therapeutic agents e.g., to compare the effects of the same agent administered via different delivery systems, or simply to assess whether a delivery vehicle itself (e.g., a viral vector or a liposome) is capable of affecting functional engraftment of a myocyte and a stem cell. These delivery vehicles may be of any form, from conventional pharmaceutical formulations, to gene delivery vehicles. For example, the therapeutic effect of the same agent administered by two or more different delivery systems (e.g., a depot formulation and a controlled release formulation), or whether a particular vehicle may have effects of itself on the tissue may be investigated.

II. Co-Cultures and Devices for use in the Methods of the Invention

The present invention also provides in vitro models of myocyte cell therapy which include a cell co-culture, comprising a base layer; a soft substrate on the base layer; a biopolymer patterned on the soft substrate; a single myocyte; and a single stem cell, wherein the single myocyte and the single stem cell are cultured such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface.

Such models, which include the foregoing cell co-culture, are suitable for use in the methods of the invention. In some embodiments, the co-cultures further include a device to measure contractility, e.g., an optical signal capturing device or a device for translating contractility into an electrical or magnetic signal, e.g., piezoresistive, piezoelectric, or strain sensor, e.g., embedded within the soft substrate.

A base layer maybe formed of a rigid or semi-rigid material, such as a metal, ceramic, or a combination thereof. In particular embodiments, the Young’s modulus of the base material used to form the base layer is greater than 1 mega-pascal (MPa). The base layer material may also be transparent, so as to facilitate observation. Examples of suitable base layer material include polymethylmethacrylate, polystyrene, polycarbonate, polyethylene terephthalate film, silicon wafer, or gold. In one embodiment, the base layer is a silicon wafer, a glass cover slip, a multi-well plate, tissue culture plate, or a chamber of a microfluidic device.

A“soft substrate” is any material that is flexible, pliable, or malleable when exposed to an external force. Other physical characteristics common to soft substrates suitable for use in the methods and devices of the invention include linear elasticity and incompressibility. Soft substrates may have a Young’s modulus in the range of about 1 to about 100,000 pascal (Pa). The soft substrates of the present invention include those that may be tunable to the stiffness of physiological tissues with a Young’s modulus of about l,000 to about 100,000 Pa, about 1,000 to about 5,000 Pa, about 1,000 to about 10,000 Pa, about 5,000 to 10,000 Pa, about 5,000 to about 20,000 Pa, about 10,000 to about 20,000 Pa, about 10,000 to about 50,000 Pa, about 20,000 to about 50,000 Pa, about 30,000 to about 50,000 Pa, about 40,000 to about 50,000 Pa, about 50,000 to about 100,000 Pa, about 60,000 to about 100,000 Pa, about 70,000 to about 100,000 Pa, about 80,000 to about 100,000 Pa, about 90,000 to about 100,000 Pa.

In some embodiments, the soft substrates have a Young’s modulus of about 0.1 Pa and about 25.0 Pa; between about 0.1 Pa and about 20 Pa; between about 0.1 Pa and about 15 Pa; between about 0.1 Pa and about 10 Pa; between about 0.1 Pa and about 5 Pa; between about 1.0 Pa and about 25.0 Pa; between about 1.0 Pa and about 20 Pa; between about 1.0 Pa and about 15 Pa; between about 1.0 Pa and about 10 Pa; between about 1.0 Pa and about 5 Pa, e.g., about 0.1, 0.5, 1.0, 1.5, 2.0, 2.5, 3.0, 3.5, 4.0, 4.5, 5.0, 5.5, 6.0, 6.5, 7.0, 7.5, 8.0, 8.5, 9.0, 9.5, 10.0, 10.5, 11.0, 11.5, 12.0, 12.5, 13.0, 13.5, 14.0, 14.5, 15.0, 15.5, 16.0, 16.5, 17.0, 17.5, 18.0, 18.5, 19.0, 19.5, 20.0, 20.5, 21.0, 21.5, 22.0, 22.5, 23.0, 23.5, 24.0, 24.5, or about 25.0 Pa. Ranges and values intermediate to the above recited ranges and values are also contemplated to be part of the invention.

Exemplary soft substrates for use in the methods and devices of the invention are described in U.S. Patent No.8,748, 181, the entire contents incorporated herein by reference. In one embodiment, the soft substrate is prepared from a single material. In another embodiment, the soft substrate is prepared from more than one material. In yet other

embodiments of the invention, a soft substrate is prepared by layering one or more soft substrates, e.g., to mimic tissue layers in vivo. Non-limiting examples of soft substrates include

polyacrylamide gels, poly(N-isopropylacrylamide), pHEMA, collagen, fibrin, gelatin, alginate, and dextran.

In one embodiment, a soft substrate is composed of polymers, such as polyacrylamide. Acrylamide may be polymerized into a gel with a finely-tuned stiffness. By varying the relative amounts of monomeric acrylamide and bis-acrylamide, the stiffness of the resulting

polyacrylamide gel may be increased (by using a higher relative amount of bis-acrylamide) or decreased (by using a lower relative amount of acrylamide). Furthermore, the addition of additives such as polypyrrole and poly-ethyl-glycol will alter the stiffness of a polyacrylamide gel.

The soft substrates can be any acrylic acid-based hydrogel constructed by free radical methacrylate). The monomeric acrylamide may be cross-linked by any diacrylate group, such as ethylegeglycol dimethacrylate and 1,4-butanediol dimethacrylate, or by N,N’

methylenebisacrylamide. The stiffness of the polymerized acrylamide may be tuned by varying the ratio of the cross-linker to the acrylamide subunit. In addition, the stiffness of the gel may be modified by co-polymerizing the acrylamide with other polymers, such as polypyrrole and poly ehtyl-glycol. The acrylamide may be co-polymerized with polyacetylene group such as polypyrrole and polyaniline to give rise to a conductive polymer.

The soft substrates may also be other soft, biocompatible gels. These include hydrogels composed of proteins such as gelatin, collagen, arginine, fibrin, and fibronectin, glucose molecules such as dextran, and glycoprotein such as hyaluronate.

Methods for preparing a soft substrate, e.g. a soft substrate comprising a biopolymer patterned on the sift substrate, include providing a base layer; depositing a sacrificial polymer on the base layer, thereby generating a sacrificial polymer layer; depositing a photoresist onto the sacrificial polymer layer, thereby generating a photoresist layer; placing a mask on top of the photoresist layer and exposing the photoresist layer to electromagnetic radiation, thereby generating a micro-patterned photoresist complementary mask; releasing the micro-patterned photoresist complementary mask from the sacrificial polymer layer; placing the micro-patterned photoresist complementary mask onto a soft substrate; and functionalizing the soft substrate by contacting the soft substrate with a linker molecule and a biopolymer. It will be understood that any suitable means for“depositing” a substance when preparing a soft substrate may be used. Depositing refers to a process of placing or applying an item or substance onto another item or substance (which may be identical to, similar to, or dissimilar to the first item or substance). Depositing may include, but is not limited to, methods of using spraying, dip casting, spin coating, 3D printing, or other methods to associate the items or substances. The term depositing includes applying the item or substance to substantially the entire surface as well as applying the item or substance to a portion of the surface.

A base layer for use in preparing a stiff substrate comprising a biopolymer patterned thereon, maybe formed of a rigid or semi-rigid material, such as a metal, ceramic, or a combination thereof. In particular embodiments, the Young’s modulus of the base material used to form the base layer is greater than 1 mega-pascal (MPa). The base layer material may also be transparent, so as to facilitate observation. Examples of suitable base layer material include polymethylmethacrylate, polystyrene, polyethylene terephthalate film, silicon wafer, or gold. In one embodiment, the base layer is a silicon wafer, a glass cover slip, a multi-well plate, tissue culture plate, or a chamber of a microfluidic device.

A sacrificial polymer layer may be applied to the rigid base layer by depositing the sacrificial polymer onto the base layer. For example, spin coating may be used to deposit the sacrificial polymer layer to the base material.“Spin coating”, as used herein, refers to a process wherein the base layer is mounted to a chuck under vacuum and is rotated to spin the base layer about its axis of symmetry and a liquid or semi-liquid substance, e.g. a polymer, is dripped onto the base layer, with the centrifugal force generated by the spin causing the liquid or semi-liquid substance to spread substantially evenly across the surface of the base layer. The resulting sacrificial polymer layer serves to temporarily secure additional coatings that are subsequently formed thereon.

In one embodiment, the sacrificial polymer is a thermally sensitive polymer that is melted or dissolved to cause the release of the micro-patterned photoresist complementary mask. An example of such a polymer is linear, non-cross-linked poly(N-Isopropylacrylamide), which is a solid when dehydrated, and which is a solid at 37^oC (wherein the polymer is hydrated but relatively hydrophobic). However, when the temperature is dropped to 32^oC or less (where the polymer is hydrated but relatively hydrophilic), the polymer becomes a liquid, thereby releasing the micro-patterned photoresist complementary mask (Feinberg et al. (2007) Science 317:1366- 1370). In another embodiment, the sacrificial polymer becomes hydrophilic, thereby releasing hydrophobic coatings, with a change in temperature. For example, the sacrificial polymer can be hydrated, crosslinked N-Isopropylacrylamide, which is hydrophobic at 37°C and hydrophilic at 32°C.

In yet another embodiment, the sacrificial polymer is an electrically actuated polymer that becomes hydrophilic upon application of an electric potential to thereby release a hydrophobic structure coated thereon. Examples of such a polymer include poly(pyrrole)s, which are relatively hydrophobic when oxidized and hydrophilic when reduced. Other examples of polymers that can be electrically actuated include poly(acetylene)s, poly(thiophene)s, poly(aniline)s,

poly(fluorene)s, poly(3- hexylthiophene), polynaphthalenes, poly(p-phenylene sulfide), and poly(para-phenylene vinylene)s.

In still another embodiment, the sacrificial polymer is a degradable biopolymer that can be dissolved to release a structure coated thereon. In one example, the polymer (e.g., polylactic acid, polyglycolic acid, poly(lactic-glycolic) acid copolymers, or nylons) undergoes time- dependent degradation by hydrolysis. In another example, the polymer undergoes time-dependent degradation by enzymatic action (e.g., fibrin degradation by plasmin, collagen degradation by collagenase, or fibronectin degradation by matrix metalloproteinase).

In yet still another embodiment, the sacrificial polymer is an ultra-hydrophobic polymer with a surface energy lower than the photoresist adhered to it. In this case, mild mechanical agitation will“pop” the micro-patterned photoresist complementary mask off. Examples of such a polymer include but are not limited to alkylsilanes (octadecyltrichiorosilane and

isobutyltrimethoxysilane), fluoroalkylsilanes (tridecafluorotetrahydrooctyltrichiorosilane, trifluoropropyltrichiorosilane and heptadecafluorotetrahydrodecyltrichlorosilane), silicones (methyihydrosiloxane - dimethylsiloxane copolymer, hydride terminated polydimethylsiloxane, trimethylsiloxy terminated polydimethylsiloxane and diacetoxymethyl terminated

polydimethylsiloxane), fluorinated polymers (polytetrafluoroethylene, perfluoroalkoxy and fluorinated ethylene propylene).

The sacrificial polymer layer provides temporary adhesion of the base layer to a photoresist layer. A photoresist layer may be applied to the sacrificial polymer layer by depositing a photoresist onto the sacrificial polymer layer. In one embodiment, spin coating (as described above with reference to applying the sacrificial polymer layer to the base layer) is used to apply a photoresist to the sacrificial polymer layer.“Photoresist” is any substance that is sensitive to electromagnetic radiation, e.g., wavelengths of light in the ultraviolet or shorter spectrum (<400 nm). A photoresist may be positive or negative. A“positive photoresist” is a type of photoresist in which the portion of the photoresist that is exposed to light becomes soluble to the photoresist developer and the portion of the photoresist that is unexposed remains insoluble to the photoresist developer.

A“negative photoresist” is a type of photoresist in which the portion of the photoresist that is exposed to light becomes essentially insoluble to the photoresist developer. The unexposed portion of the photoresist is solubilized by the photoresist developer.

Non-limiting examples of photoresist materials include 1-methoxy-2-propanol acetate (SU-8), bisazides, poly(vinyl cinnamate), and novolaks, polymethylmethacrylates (PMMA), epoxies.

A micro-patterned photoresist complementary mask comprises the photoresist and the sacrificial polymer layer, a mask (comprising a desired shape), e.g., a solid mask, such as a photolithographic mask, which is provided and placed on top of the photoresist layer.

Subsequently, a portion of the photoresist layer (i.e., the portion of the photoresist not covered by the solid mask) is exposed to electromagnetic radiation. A suitable shape may be any desired shape, such as a geometric shape, e.g., a line a circle, square, rectangle, triangle, line, or combinations thereof. In other embodiments, the shape may be the shape of an organ, or portion thereof.

The mask placed on top of the photoresist layer is typically fabricated by standard photolithographic procedure, e.g., by means of electron beam lithography. Other methods for creating such masks include focused energy for ablation (micromachining) including lasers, electron beams and focused ion beams. Similarly, chemical etchants may be used to erode materials through the photoresist when using an alternative mask material. Examples of chemical etchants include hydrofluoric acid and hydrochloric acid.

Any suitable material, e.g., a material that has a flat surface, e.g., a metal (gold, silver, platinum, or aluminum), a ceramic (alumina, titanium oxide, silica, or silicon nitride), may be used for making the mask.

In order to release the sacrificial polymer layer from the micro-patterned photoresist complementary mask, the sacrificial polymer layer is exposed to conditions that dissolve or allow the sacrificial polymer layer to switch states.

For example, a sacrificial polymer layer formed of poly-N-iso-propylacrylamide

(PIPAAM) (non-cross-linked) will dissolve in an aqueous media at a temperature less than 32°C. In another example, a sacrificial polymer layer is formed of PIPAAM (cross-linked) will switch from a hydrophobic to hydrophilic state in an aqueous media at a temperature less than 32°C. The hydrophilic state will release the sacrificial polymer layer from the micro-patterned photoresist complementary mask. In yet another embodiment, the sacrificial polymer layer includes a conducting polymer, such as polypyrrole, that can be switched from a hydrophobic to hydrophilic state by applying a positive bias that switches the conducting polymer from a reduced to an oxidized state. In additional embodiments, the sacrificial polymer layer can include a degradable polymer and/or polymer that undergoes time-dependent degradation by hydrolysis (as is the case, for example, for polylactic and polyglycolic acid) or by enzymatic action (for example, fibrin degradation by plasmin).

Once the micro-patterned photoresist complementary mask is released from the sacrificial polymer layer, it is place onto a soft substrate and the soft substrate is functionalized as described above.

In order to functionalize a soft substrate, a soft substrate is contacted with a functional linker molecule, e.g., a bifunctional or multifunctional linker molecule, and a biopolymer. The patterned photoresist complementary mask may be removed from the soft substrate prior to the addition of the biopolymer or may remain in place while the soft substrate is contacted with the biopolymer. In one embodiment, following contacting the soft substrate with a linker molecule, e.g., a bifunctional linker molecule, a second patterned photoresist complementary mask is placed in conformal contact with the linker molecule deposited soft substrate.

A“linker molecule”, as used herein, is well-known in the art and refers to a first molecule that may associate with a second molecule to attach the second molecule to a third molecule, where the first, second, and third molecules may be identical, similar, or dissimilar to one another. These may include bifunctional linker molecules and multifunctional linker molecules. A bifunctional linker molecule has two linking moieties, e.g., two linking moieties that are activated using different methods. A multifunctional linker molecular has at least three linking moieties, e.g., where the three linking moieties include at least two linking moieties that are activated using different methods. Non-limiting examples of a functional linker molecules include N- Sulfosuccinimidyl 6-hexanoate (sulfo-SANPAH) or N-succinimidyl ester of acrylamidohexanoic acid.

A linker molecule may be activated prior to contacting the soft substrate with an appropriate biopolymer.“Activating”, e.g., activating a linking molecule, refers to use of a method that allows a linking moiety of a linking molecule to associate with another molecule. Non-limiting examples of activating methods include ultraviolet radiation, chemical activation, thermal, or physical activation. For example, sulfo-SANPAH contains a nitrophenyl azide, which is activated by ultraviolet light, and a sulfo-N-hydroxysuccinimide ester, which is chemically activated by amino groups.

“Biopolymer” refers to any proteins, carbohydrates, lipids, nucleic acids or combinations thereof, such as glycoproteins, glycolipids, or proteolipids.

Examples of suitable biopolymers that may be used for substrate functionalization include, without limitation:

(a) extracellular matrix proteins to direct cell adhesion and function (e.g., collagen, fibronectin, laminin, vitronectin, or polypeptides (containing, for example the well known -RGD- amino acid sequence));

(b) growth factors to direct specific cell type development cell (e.g., nerve growth factor, bone morphogenic proteins, or vascular endothelial growth factor);

(c) lipids, fatty acids and steroids (e.g., glycerides, non-glycerides, saturated and unsaturated fatty acids, cholesterol, corticosteroids, or sex steroids);

(d) sugars and other biologically active carbohydrates (e.g., monosaccharides, oligosaccharides, sucrose, glucose, or glycogen);

(e) combinations of carbohydrates, lipids and/or proteins, such as proteoglycans (protein cores with attached side chains of chondroitin sulfate, dermatan sulfate, heparin, heparan sulfate, and/or keratan sulfate); glycoproteins (selectins, immunoglobulins, hormones such as human chorionic gonadotropin, Alpha fetoprotein or Erythropoietin (EPO)); proteolipids (e.g., N- myristoylated, palmitoylated and prenylated proteins); and glycolipids (e.g., glycoglycerolipids, glycosphingolipids, or glycophosphatidylinositols);

(f) biologically-derived homopolymers, such as polylactic and polyglycolic acids and poly-L-lysine;

(g) nucleic acids (e.g., DNA or RNA);

(h) hormones (e.g., anabolic steroids, sex hormones, insulin, or angiotensin);

(i) enzymes (e.g., oxidoreductases, transferases, hydrolases, lyases, isomerases, ligases; examples: trypsin, collegenases, or matrix metalloproteinases);

(j) pharmaceuticals (e.g., beta blockers, vasodilators, vasoconstrictors, pain relievers, gene therapy, viral vectors, or anti-inflammatories);

(k) cell surface ligands and receptors (e.g., integrins, selectins, or cadherins); and (1) cytoskeletal filaments and/or motor proteins (e.g., intermediate filaments,

microtubules, actin filaments, dynein, kinesin, or myosin).

Typically, a soft substrate will be contacted with a single linker molecule and a single biopolymer. However, a soft substrate may be contacted with two or more linker molecules and/or two or more biopolymers to create, e.g., a boundary condition, such as those described in PCT/US2008/011173, the contents of which are incorporated in their entirety by reference. For example, one linker molecule may be used to adhere a particular biopolymer to the soft substrate, e.g., a biopolymer with a particular cell type specificity, and a second linker molecule may be used to adhere a particular biopolymer with a different cell type specificity to the soft substrate. In practicing the invention, a variety of techniques can be used to foster selective cell adhesion of two or more cell types to the substrate. Included, without limitation, are methods such as localized protein adsorption, organosilane surface modification, alkane thiol self-assembled monolayer surface modification, wet and dry etching techniques for creating three-dimensional substrates, radiofrequency modification, and ion-implantation (Lom et al., 1993, J. Neurosci. Methods 50:385-397; Brittland et al., 1992, Biotechnology Progress 8:155-160; Singhvi et al., 1994, Science 264:696-698; Singhvi et al., 1994, Biotechnology and Bioengineering 43:764-771; Ranieri et al., 1994, Intl. J. Devel. Neurosci.12(8):725-735; Bellamkonda et al., 1994,

Biotechnology and Bioengineering 43:543-554; and Valentini et al., 1993, J. Biomaterials Science Polymer Edition 5(1/2):13-36).

In another embodiment, the soft substrates are functionalized by contacting the soft substrate with a repellent. As used herein, a "repellent" is a composition that, relative to the substrate to which it is applied, inhibits adhesion of a biopolymer and/or particular cell type, thereby causing a first particular cell type to adhere preferentially to the substrate. Agarose, hyaluronic acid, and alginate are examples of suitable repellents. In this embodiment, the patterned photoresist complementary mask is removed (or in the case of use of multiple patterned photoresist complementary masks, the first patterned photoresist complementary mask is removed and optionally a second patterned photoresist complementary mask is placed on the soft substrate) prior to, e.g., seeding cells.

In one embodiment, the biopolymer is an extracellular matrix (ECM) protein, e.g., a fibronectin, a laminin, a collagen, a gelatin, a fibrin, a vitronectin, or a fibrinogen, or

combinations thereof. The biopolymer is deposited on the soft substrate in a pattern that allows the cells to align substantially anisotropically along their length to the biopolymer patterned on the soft substrate, In one embodiment, the biopolymer is patterned spatially using alternating high-density lines of biopolymer with either low-density biopolymer or a chemical that prevents protein adhesion (e.g., Pluronics F127). The dimensions of these lines are typically 10-100 µm width at 2-100 µm spacing, (see, e.g., Feinberg, Science 317:1366 - 1370, 2007), however, the width and spacing may be altered to change the alignment. Changes in alignment in turn affect the structural and functional anisotropy and anisotropy of cells and tissues. The width and spacing of the ECM lines may be varied over the range from about 0.1 µm to about 1000 µm, from about 1 µm to about 500 µm, from about 1 µm to 250 µm, from about 1 µm to 100 µm, from about 1 µm to 90 µm, from about 1 µm to 80 µm, from about 1 µm to 70 µm, from about 1 µm to 60 µm, from about 1 µm to 50 µm, from about 1 µm to 40 µm, from about 1 µm to 30 µm, from about 1 µm to 20 µm, from about 1 µm to 10 µm, from about 2 µm to 100 µm, from about 2 µm to 90 µm, from about 2 µm to 80 µm, from about 2 µm to 70 µm, from about 2 µm to 60 µm, from about 2 µm to 50 µm, from about 2 µm to 40 µm, from about 2 µm to 30 µm, from about 2 µm to 20 µm, from about 2 µm to 10 µm, from about 1 µm to 100 µm, from about 5 µm to about 100 µm, from about 5 µm to about 90 µm, from about 5 µm to about 80 µm, from about 5 µm to about 70 µm, from about 5 µm to about 60 µm, from about 5 µm to about 50 µm, from about 5 µm to about 40 µm, from about 5 µm to about 30 µm, from about 5 µm to about 20 µm, and from about 5 µm to about 20 µm. Values and ranges intermediate to the above-recited values and ranges are also contemplated by the present invention.

The width and spacing of the ECM lines can be equivalent or disparate. For example, both the width and spacing can be about 0.1, about 0.2, about 0.25, about 5, about 0.75, about 1, about 2, about 3, about 4, about 5, about 6, about 7, about 8, about 9, about 10, about 11, about 12, about 13, about 14, about 5, about 16, about 17, about 18, about 19, or about 20 µm. In other embodiments, the width can be about 0.1, about 0.2, about 0.25, about 5, about 0.75, about 1, about 2, about 3, about 4, about 5, about 6, about 7, about 8, about 9, or about 10 µm and the spacing can be about 1, about 2, about 3, about 4, about 5, about 6, about 7, about 8, about 9, about 10, about 11, about 12, about 13, about 14, about 5, about 16, about 17, about 18, about 19, about 20, about 21, about 22, about 23, about 24, about 25, about 26, about 27, about 28, about 29, about 30, about 31, about 32, about 33, about 34, about 35, about 36, about 37, about 38, about 39, or about 40 µm. Conversely, the width can be about 1, about 2, about 3, about 4, about 5, about 6, about 7, about 8, about 9, about 10, about 11, about 12, about 13, about 14, about 5, about 16, about 17, about 18, about 19, about 20, about 21, about 22, about 23, about 24, about 25, about 26, about 27, about 28, about 29, about 30, about 31, about 32, about 33, about 34, about 35, about 36, about 37, about 38, about 39, or about 40 µm and the spacing can be about 0.1, about 0.2, about 0.25, about 5, about 0.75, about 1, about 2, about 3, about 4, about 5, about 6, about 7, about 8, about 9, or about 10 µm. Values and ranges intermediate to the above-recited values and ranges are also contemplated by the present invention.

In one embodiment, e.g., the width of the biopolymer lines is about 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, or about 25 µm and the spacing of the lines is about 0.5, 1, 1.5, 2, 2.5, or about 3 µm. Values and ranges intermediate to the above-recited values and ranges are also contemplated by the present invention.

Typically the patterned biopolymer lines are parallel to one another, but they may also be at angles to one another.

To seed cells, soft substrates are placed in culture with a cell suspension allowing the cells to settle and adhere to the soft substrate. In the case of an adhesive surface treatment, cells bind to the material in a manner dictated by the surface chemistry. The cells on the substrates may be cultured in an incubator under physiologic conditions (e.g., at 37°C).

This invention is further illustrated by the following examples which should not be construed as limiting. The contents of all references, patents and published patent applications cited throughout this application, as well as the Figures, are hereby incorporated by reference.

EXAMPLES

Materials and Methods

The following materials and methods were used in the present invention.

Cell Culture

Cor.At mES- and miPS-derived cardiac myocytes

Cor.At mES- (Lot# CS25CL_V_SN_1M, production date: 2010-02-16; CS148P_1M, production date: 2010-07-09; and CS152_1M, production date: 2010-08-16) and miPS- (Lot# CS4i4M, (AXIO0013_1VL); CS07CL-i, production date: 2011-05-09) derived cardiac myocytes were cultured using culture medium prepared by and according to manufacture specifications (Axiogenesis, Cologne, Germany). Briefly, cells were cultured in culture flasks pre-coated with 10 µg/ml fibronectin in puromycin-containing culture media at 37°C and 5% CO₂ for 24 hrs. After 48 hours, the cells were rinsed with fresh media, without the puromycin antibiotic selection agent.

Harvest of neonatal mouse cardiomyocytes

Ventricular tissue was excised from 2-day old neonatal Balb/c mice, and dissociated into a 95% pure preparation of isolated cardiac myocytes as previously described (Sheehy et al., 2014 Stem Cell Reports.2:282-294). Excised ventricular tissue was rinsed with Hanks balanced salt solution, incubated in a 0.1% (w/v) solution of trypsin (USB Corp., Cleveland, OH) with agitation overnight at 4°C, and dissociated via serial exposure to a 0.1% (w/v) solution of collagenase type II (Worthington Biochemical, Lakewood, NJ) at 37ºC for 2 minutes. A 45 minute pre-plating step was used to reduce the number of non-myocytes in the preparation. All procedures were approved by the Harvard animal care and use committee.

Primary and stem cell-derived cardiomyocyte seeding procedures

Cor.At mES- and miPS-derived cardiac myocytes were cultured, according to

manufacture specifications (Axiogenesis, Cologne, Germany), in a positive selection media containing puromycin for 48 hours before cell dissociation by 0.25% trypsin. For heterogeneous cultures, Cor.At mES- or miPS-CMs were seeded simultaneously with isolated neonatal mouse ventricular myocytes onto microcontact printed substrates at equal densities (8,000cells/cm² per cell type) and cultured in Cor.At proprietary medium (Axiogenesis, Cologne, Germany). For homogeneous cultures, primary myocytes or mES-.miPS-CMs were seeded at densities of 16,000 cells/cm². Myocytes were cultured on microcontact printed substrates for three days before experimentation. Experiments were carried out in an incubation chamber on a Zeiss laser scanning confocal microscope maintained at 37°C and cultured in Tyrode’s solution (1.8 mM CaCl₂, 5 mM glucose, 5 mM HEPES, 1 mM MgCl₂, 5.4 mM KCl, 135 mM NaCl, 0.33 mM NaH₂PO₄, pH 7.4).

Photolithography and micro-contact printing

Polymer stamps designed for micro-contact printing were prepared as previously described (McCain et al., 2012b, Proc Natl Acad Sci U S A.109:9881-9886; McCain et al., 2014, Am J Physiol Heart Circ Physiol.306:H1525-1539). Photolithographic masks were designed in AutoCAD (Autodesk Inc., San Rafael, CA), and consisted of rectangles of 2000 µm² surface area, spaced by 100 µm on each side, with length:width ratios of 7:1. Silicon wafers (Wafer World, West Palm Beach, FL) were spin-coated with SU-82002 photoresist (MicroChem Corp, Newton, MA), exposed to UV light, and submerged in propylene glycol methyl ether acetate to dissolve masked regions. Polydimethylsiloxane (PDMS, Sylgard 184, Dow Corning, Midland, MI) elastomer was poured onto the wafers, cured, and peeled to generate stamps.

Fabrication and microcontact printing of polyacrylamide gels

All functional experiments were conducted using 13 kPa or 90 kPa polyacrylamide gel substrates (5% acrylamide:0.3% bisacrylamide) (Molecular Probes, Eugene, OR, USA) that were fabricated on 25 mm coverslips, as previously described (McCain et al., 2012, Am J Physiol Heart Circ Physiol.302:H443-450; McCain et al., 2014, Am J Physiol Heart Circ Physiol. 306:H1525-1539; Ye et al., 2014, Integr Biol (Camb).6:152-163). For Traction Force

Microscopy experiments, gels were doped with streptavidin-acrylamide and 200 nm fluorescent beads (Invitrogen) with a final ratio of 1:5 and 1:30 by volume, respectively. For all other experiments, gels were bead-free. Fibronectin (BD Biosciences, Bedford, MA) was biotinylated by cross-linking with biotin using Sulfo-NHS-LC-Biotin (Pierce, Rockford, IL). PDMS stamps were incubated with 200 µg/mL biotinylated FN for 1hr and dried before stamping the top surface of polyacrylamide gels.

Immunofluorescence

Myocytes were fixed in 4% PFA in PBS buffer at room temperature for 10 minutes. Cells were immunostained by incubating for 1hr at room temperature with combinations of primary antibodies against Cx43 (MAB3068; Millipore), β-catenin (C2206; Sigma-Aldrich), sarcomeric α-actinin (clone EA-53; Sigma-Aldrich), vinculin (hVIN-1; Sigma-Aldrich), 4',6'-diamidino-2- phenylindole hydrochloride (DAPI; Invitrogen), and Alexa Fluor 488-, 546-, or 633-conjugated Phalloidin (Invitrogen). Secondary antibodies conjugated to Alexa Fluor 488, 546, or 633 (Invitrogen) were used against the appropriate primary antibody for 1hr incubation at room temperature. All antibodies were used at a dilution of 1:200. Immunostained samples were imaged with a Zeiss LSM 5 LIVE confocal microscope. Traction Force Microscopy (TFM)

Spontaneously beating cardiac myocytes along with 200 nm fluorescent beads at the top surface of polyacrylamide gels were imaged sequentially at 60 Hz using a Zeiss LSM 5 LIVE scanning confocal microscope, with a 40X Plan-Apochromat oil objective. Cells were imaged at 37°C for at least four contraction cycles. High-resolution traction force microscopy Fourier transform traction cytometry methods were used to track bead displacement over time and to determine traction stress as previously described (Butler et al., 2002, Am J Physiol Cell Physiol. 282:C595-605). The total traction force, F_x, exerted by a cell on the substrate along the x-axis was calculated by summing the magnitudes of all traction force vectors, and multiplying by a factor of one half,

Ratiometric Ca²⁺ experiments and analysis

Myocytes were incubated with in a 8 µM solution of acetoxymethyl (AM) Fura Red (Invitrogen, F-3021) reconstituted in Pluronic F-127 (20% solution in DMSO; Invitrogen, P- 3000MP) for 20 minutes before rinses in Tyrode’s solution and experimentation. Spontaneously beating cardiac myocytes were imaged sequentially at 216.5 Hz using a Zeiss LSM 5 LIVE scanning confocal microscope, with a 40X Plan-Apochromat oil objective, at 37C. Images were collected at 405 nm and 488 nm and pre-processed using ImageJ software to correct background illumination and to outline each cell within a pair (GFP positive mES- and miPS-CMs were distinguishable from neonate myocytes). The mean intensity over time in each ROI was measured and analyzed in MATLAB. ROI measurements were parsed to isolate the time course of the Ca²⁺ transients, as measured under 405 nm (Ca405) and 488 nm (Ca488) illumination, and utilized to calculate the ratio Ca405/Ca488. Only traces pertaining to cells beating at a frequency comparable with those observed during TFM experiments (2.5 ± 0.8 Hz) were included in this phase.4-6 steady states transients were averaged in each recording and utilized to calculate diastolic level, peak level, time to peak and the duration of the Ca²⁺ transient at 50% and 90% decay. Two separate analyses of the ratiometric recording were performed. First, the trains of calcium transients recorded in each cell in the pair were subjected to cross correlation analysis (Cohen-Karni et al., 2009). In brief, the normalized cross-correlation of two signal x and y was calculated with the formula

where are the sampled values of the signals; are the means and

standard deviations of x and y respectively and represents a time lag. The value of for which is maximum is an estimates of the conduction velocities, as it determines how many frames it takes for the peak of one signal to travel to the second one. The maximum value of (or the cross-correlation coefficient) provides an estimate of how globally similar the two signals are. Statistical Analysis

Statistical analysis was conducted with SigmaPlot: data were first tested with the Shapiro- Wilk test for normality and the Levene median test for equal variance.1-way ANOVA tests were conducted for normally distributed datasets, followed by pairwise post-hoc Student–Newman– Keuls test. Alternatively, statistical conclusions were drawn using ANOVA on ranks followed by Dunn’s test for pairwise comparison.

Cell Mechanics Model

In the following finite element model, the cell-pair µtissue is treated as a two-dimensional (2D) continuum (Figure 7A). The cell-substrate adhesion is modeled by applying elastic constraints to the displacement field throughout the micro-tissue domain (Figure 7B). Thus, the equilibrium equations of the 2D µtissue in plane can be written as (Sadd, 2014, Elasticity, Third Edition: Theory, Applications, and Numerics)

where , , are stress components, and are displacement components, is the height of the µtissue, and the cell-substrate linkage constant. The traction stress components

on the substrate can be written as

The magnitude of the traction stress was plotted for the traction stress

maps in Figure 4 in the text and in Figure 7. Note that in the 2D problem the cell-substrate adhesion enters the equilibrium equations as“body force”, instead of boundary conditions. The position-dependent cell-substrate linkage constant

is a lumped parameter that accounts for both the substrate elasticity and the density of bound integrin. Because the substrate elasticity is not varied in this work,

is defined as a function of the density of bound integrin

alone

where parameter is the maximal cell-substrate linkage stiffness and parameter

is the maximal density of bound integrin.

were used in all simulations. The focal adhesion maturation is thought to be coupled with the traction stress with a positive feedback loop (Grosberg et al., 2011, PLoS Comput Biol.7:e1001088). Here the positive feedback was implemented using the following simple form

where is the traction stress threshold, referencing to which the density of bound integrin is redistributed based on the magnitude of the traction stress.

a was used for the homogeneous pair and

for the heterogeneous pair in the simulations. The purpose of applying Eq. (6) is to make the bound integrin density

approaching

when

and to make it approaching zero when

The cell-cell adhesion is modeled as elastic springs and implemented into the finite element model by defining its deformation energy

where is the cell-cell adhesion spring constant, the path denotes the cell-cell interface (Figure 4A in the text), denotes arch length along are displacement

components of cell 1 and cell 2 at the cell-cell interface, respectively. The equation

kPa was used for the homogeneous pair and for the heterogeneous pair in the

simulations. Furthermore, Hill’s three-element model (Fung, 1993, Biomechanics: Mechanical Properties of Living Tissues) was adopted for the contraction of cardiomyocytes. A schematic of Hill’s three-element model is shown in Figures 7B-7C, in which the myofibril is modeled as a contractile element and an elastic spring in series, and a parallel element represents the passive components of the cytoskeleton that deforms under active contraction. The stress-strain equations of the three-element model take the following forms when the direction of the myofibril is parallel to the x-axis of the coordinate system,

where Eq. (8) represents the overall stress-strain relation of the three-element model shown in Figures 7B-7E and are elastic moduli of the parallel element, are strain

components, is the contractile stress. Equation (9) is the linearized relation for the contractile element, and and are the isometric tension and the strain rate constant of the contractile element, respectively. Equation (10) simply represents the strain summation for the series element and contractile element, and is the elastic modulus of the series element. Note that the

contractile element only generates tensile stress along the direction of myofibril. Thus, the myofibril orientation needs to be specified in the simulations. In all simulations, we assume the myofibrils of the µtissue align parallel to the longitudinal axis. A list of all parameters in reported in Table 1.

Table 1. Summary of the parameters and variables used in the finite element simulations.

Finite element simulations

Equations (1)-(9) were implemented in a finite element program. The cell-pair µtissue is meshed using 3-node triangular elements with mesh size of ~2.0 µm. The total area of the µtissue is 2500 µm² and the aspect ratio is 7:1. The thickness of the µtissue is

The finite element domain is divided into two regions: cell 1 (gray region) and cell 2 (white region), as shown in Figure 4A in the text. Finite element nodes of two different cells on the cell-cell interface overlap and are connected by elastic springs derived from Eq. (7). The finite element model is focused on the time period of diastolic to peak systolic contraction and is a dynamics process. Explicit time integration scheme was used to obtain time- dependent solutions of stress and strain fields from diastolic to peak systolic contraction, in which the time increment is set to be one millisecond, and the time to peak systolic contraction is set to be 80 milliseconds for all simulations. For the initial conditions of the simulations, the stress and strain components are set to be zero at time =0 at the diastolic state.

Simulations were first performed to determine the distribution of bound integrin in the steady state. Assuming the uniform distribution of bound integrin (i.e., = 400 µm^-2 on the entire island) at the beginning, traction stress was calculated at the peak systolic contraction. The magnitude of the traction stress was then substituted into Eq. (6) to update the distribution of bound integrin. The updated distribution of bound integrin was substituted into Eq. (5) to calculate the cell-substrate linkage constant at the steady state. In the present model, we are more concerned about the qualitative aspect of the problem, thus only one iteration was run to obtain the steady state. This procedure was performed for the homogeneous (Case 1) and heterogeneous (Case 2) cell pairs. The cell-substrate linkage constant at the steady state was then used in the new simulations in which the final traction stress, cellular tension (i.e., ), and shortening were predicted, which are presented in Figures 4B-4D in the text.

Results

Contractile Structure and Function in Primary and Stem Cell-Derived Myocytes

In this study, myocytes harvested from neonate murine hearts or differentiated from mES and miPS cells were used to model stronger native and weaker regenerated myocardium, respectively. To validate this choice, the structural and functional proficiency of isolated neonate, mES- and miPS-CMs cultured on fibronectin islands (7:1 length to width ratio) microcontact printed on soft (13 kPa) gels that mimic the microenvironment of the healthy heart (Engler et al., 2008, J Cell Sci.121:3794-3802; McCain et al., 2014, Am J Physiol Heart Circ Physiol.306:H1525-1539) were assessed.

Neonate, mES- and miPS-CMs exhibited striated myofibrils that extended parallel to the longitudinal axis of the cell, as demonstrated by immunostains of actin (Figure 1Ai) and α-actinin (Figure 1Aii). To quantify actin alignment, the orientational order parameter (OOP), which yields values ranging from 0 to 1 for randomly distributed and perfectly aligned networks, respectively (Pasqualini et al., 2015, Stem Cell Reports.4:340-347) was calculated. It was found that both primary and stem cell-derived myocytes had highly aligned cytoskeleton with OOP>0.9 (Figure 1B). To compare the contractile force of spontaneously beating myocytes, traction force microscopy was used. Substrates were doped with fluorescent beads whose displacement between peak systole and diastole was measured and converted into traction stress (Butler et al., 2002, Am J Physiol Cell Physiol.282:C595-605). Displacement (Figure 1Ci) and traction stress (Figure 1Cii) heat maps of neonate, mES- and miPS- CMs revealed localization of stresses at the proximal ends of the cells, with large traction force vectors along the longitudinal axis. A comparison of peak systolic longitudinal force revealed roughly two-fold lower magnitudes for mES- and miPS-CMs relative to neonate myocytes (Figure 1D). Taken together these results show that despite exhibiting similar cytoskeletal organization, stem cell-derived myocytes were significantly weaker than neonate myocytes.

Contractile Architecture in Paired Primary and Stem Cell-Derived Myocytes

For cardiac stem cell therapy to be effective, seamless integration of newly formed and existing myocardium must be achieved. This requires the formation of intercalated disks that facilitate gap junction formation and electrical conduction (Chung et al., 2007, J Cardiovasc Electrophysiol.18:1323-1329; Kim et al., 2010, Proc Natl Acad Sci U S A.107:565-570), as well as myofibril registration and force transmission (Parker and Ingber, 2007, Philos Trans R Soc Lond B Biol Sci.362:1267-1279). To determinewhether this process is impaired when cells of unequal contractile strength are coupled together, cardiac myocyte pairs, or µtissues (McCain et al., 2012a, Am J Physiol Heart Circ Physiol.302:H443-450; McCain et al., 2012b, Proc Natl Acad Sci U S A.109:9881-9886) were engineered, Homogeneous µtissues consisted solely of pairs of neonate, mES- or miPS-CMs; while heterogeneous µtissues were composed of a single neonate cell coupled to either a GFP⁺ mES- or miPS-CM (Figure 2A). It was observed that myofibrils aligned parallel to the longitudinal axis and appeared registered in all pairs (Figure 2A). All µtissue configurations formed cell-cell junctions and expressed β-catenin (Figure 2B) and connexin-43 (Cx43) (Fig 2C), proteins responsible for mechanical and electrical coupling, respectively. Quantitatively, myocytes spread to occupy roughly half the µtissue area, exhibited actin OOP >0.9, sigmoidal junctions (McCain et al., 2012b, Proc Natl Acad Sci U S A.

109:9881-9886) and elongated nuclei (Ye et al., 2014, Integr Biol (Camb).6:152-163) independently of pairing type (Figure 5). Together, these results demonstrate that in our cardiac cell therapy on-a-chip platform, heterogeneous µtissues can exhibit the same molecular markers of engraftment that are observed in vivo. Calcium Handling in Paired Primary and Stem Cell-Derived Myocytes

To test the electro-chemical coupling of engineered µtissues, Ca²⁺ transients via dual- excitation ratiometric analysis (Lohr, 2003, Cell Calcium.34:295-303; Sheehy et al., 2014, Stem Cell Reports.2:282-294) were quantified. While Ca²⁺ transients in neonate, mES- and miPS-CM paired in heterogeneous pairs had different time courses, all coupled cells exhibited synchronized activity (Figure 2D). A cross-correlation function that measures the time-lag between the two Ca²⁺ signals was constructed and, within the limits of the experimental sampling frequency, no frame lag was observed (Figure 2Ei). This translates to a conduction velocity in excess of 5 cm/s, in good agreement with previous reports (Cohen-Karni et al., 2009, Proc Natl Acad Sci U S A. 106:7309-7313). Using the cross-correlation coefficient, a metric of similarity between signals, it was demonstrated that the time course of cytoplasmic Ca²⁺ in homogeneous mES pairs, and all heterogeneous pairs, had significantly more variability than the other homogeneous pairs (Figure 2Eii). In addition, while homogeneous pairs maintained nearly identical diastolic Ca²⁺ levels, mES-CMs had almost two-fold lower Ca²⁺ levels than neonate myocytes and 1.5 times lower levels than miPS-CMs. Moreover, diastolic Ca²⁺ levels in neonate myocytes paired with mES- or miPS-CMs were ~30% lower than that observed in homogeneous neonate pairs. Correspondingly, the coupling of stem cell-derived myocytes to neonate myocytes did not increase diastolic Ca²⁺ levels; mES-CMs exhibited no significant change while miPS-CMs experienced a significant ~9% reduction when compared to respective homogeneous pairings (Figure 2F).

Taken together, these results demonstrate that myocytes at different maturation stages can form synchronized µtissues, yet possible cell line-specific pro-arrhythmic heterogeneities in Ca²⁺ handling persist and contribute to depressed contractile function, perhaps through a classical inotropic effect (Bers, 2001, Nature.415:198-205).

Contractile Function in Paired Primary and Stem Cell-Derived Myocytes

To test if intrinsic differences in force generation capacity between primary and stem cell- derived myocytes impact µtissue function, traction force microscopy was used to study spontaneously beating cell pairs. A comparison of substrate displacement (Figure 3Ai) and traction stress (Figure 3Aii) across homogeneous µtissues revealed that paired neonatal myocytes exerted greater peak systolic displacement and traction stress than either mES- and miPS-CM µtissues. Importantly, traction stresses adjacent to the cell-cell junctions were evident in heterogeneous pairs, but virtually absent in homogeneous ones. Confocal imaging of a focal adhesion marker, vinculin, revealed focal adhesion formation at the lateral ends of all cell-pair types and occasional localization of adhesions near heterogeneous cell-cell junctions, consistent with regions of high traction stress (Figure 3B).

To assess whether neighboring neonate, mES or miPS-CMs exhibit mechanical synchrony when coupled, the total longitudinal traction force exerted on the substrate in each cell, and at the junction during a contraction cycle was quantified (Figure 3Ci). Both mES- and miPS- CMs remained weaker than neonate myocytes in homogeneous pairs, but exhibited synchronous contraction profiles and minimal junctional traction force (Figure 3Cii). In heterogeneous pairs, contraction profiles for neonate, mES- or miPS-CMs were also in-phase (Figure 3ACiii-iv), yet the total durations of the contraction cycles were ~ 40% longer than in homogeneous neonate pairs (Figure 3D). Furthermore, in homogeneous pairs, neighboring cells exerted similar levels of force on the substrate: primary myocytes exerted significantly higher levels of force (~0.55µN, Figure 3E) than mES- and miPS-CMs (~0.25 µN and ~0.30 µN, respectively). In contrast, the difference in forces between neighboring cells within heterogeneous pairs comprised 30-35% of the total peak systolic force exerted by the cell pair. Finally, it was determined whether the variability in tension generated by cell pairs resulted from differences in the performance of each cell within the pair, which might be actively contracting or passively stretching. In homogeneous pairs of primary myocytes, each cell within a pair shortened by roughly 7% (Figure 3F), significantly more than in the stem cell-based homogeneous pairs (3% and 4% for mES- and miPS-CMs, respectively). In heterogeneous pairs, mES- and miPS-CMs shortened by 1.6% and 4.6%, respectively, while neonate myocytes within these pairs shortened by 5.6% and 7.7%, respectively. These values resulted predominantly from movement by lateral edges in the direction of the neighboring cell, suggesting both myocytes were actively contracting, and not passively stretching.

These results suggest that the cell-matrix adhesions located at the cell-cell junctions may have the functional role of dissipating excess force onto the substrate thereby preventing mechanical disruption of the cell-cell junction.

A Computational Model of Paired Primary and Stem Cell-Derived Myocyte Mechanics

Multiple transcriptional and functional differences that exist between these primary and stem cell-derived myocyte populations, including expression of atrial transcription factors, fetal contractile protein isoforms, and reduced calcium currents have been documented (Sheehy et al., 2014. Stem Cell Reports.2:282-294). Since the relative importance of each mechanism may vary in newly formed myocytes from different species, origins, and/or repair mechanisms, a generalized computational model of µtissue mechanics was developed (Figure 4A). Through this model, it was directly tested whether differences in contractile strength may be responsible for the pattern of adhesion formation and traction stress observed in this study. The µtissue was treated as a two dimensional continuum whose contractile behavior is governed by Hill’s equation and whose anchoring is mediated by elastic constraints. A positive feedback between adhesion maturation and traction stress was additionally included (Grosberg et al., 2011. PLoS Comput Biol.7:e1001088). Model parameters based on literature values and our single cell data were assigned, exclusively. Specifically, smaller magnitudes of isometric tension and strain rate constants were assigned to stem cell-derived myocytes in heterogeneous pairs, due to the known inotropic effect of diastolic calcium levels (Bers, 2001. Nature.415:198-205). Importantly, the locations of adhesion formation were not specified, but allowed for adhesion maturation and strengthening to develop in the model (Figure 7).

The simulations showed that in homogeneous µtissues, high traction stress was distributed solely near the two lateral ends of the islands. In heterogeneous µtissues, additional high traction stress localized to the tip of the neonate myocyte near the cell-cell interface (Figure 4B, arrow). These results were consistent with the experimental traction stress maps (Figure 3A), demonstrating that the presence of a weaker cell in the pair was sufficient to cause the patterns of traction stress observed in heterogeneous cell pairs. Further, as shown in Figure 4C, the predicted cellular tension at peak systolic contraction was distributed continuously across the cell-cell interface for the homogeneous pair. In contrast, in the heterogeneous pair, cellular stress was lower in the stem cell-derived myocyte than in the neonate one, and a discontinuity of cellular stress occurred at the cell-cell interface. Moreover, predicted forces and the average percent shortening (Figure 4D) were also in agreement with the in vitro data (Figures 3E-F), confirming that the stem cell-derived myocytes shortened less than the neonate myocyte, yet neither cell lengthened.

Thus, this computational model suggests that pairing myocytes with intrinsically different contractile properties, such as a reduction in isometric tension, is sufficient to induce force transmission remodeling at the cell-cell junction.

Mechanotransduction and Cardiac Stem Cell Therapy

Using the in vitro devices and methods described herein, the structural and functional engraftment of primary and stem cell-derived myocytes have been qualified and quantified. It has been demonstrated that weaker mES- or miPS-CMs can align myofibrils, form gap junctions, and contract synchronously with stronger neonate myocytes. Further, it has been shown that stem cell-derived myocytes exhibit impaired calcium handling and exert lower contractile force than primary cells, resulting in a net junctional force that was balanced by compensatory cell-substrate adhesions at the cell-cell junction. The in vitro and silico results described herein demonstrate that myocytes sense the force imbalance across the cell-cell junction and respond depositing junctional substrate adhesions that dissipate the excess force. Together these characteristics demonstrate that newly formed myocytes are weaker than spared ones in vivo, and therefore subjected to the compensatory mechanism described herein. In this context, the data demonstrate that stem cell-derived myocytes can form a syncytium with the native myocardium, which may explain the reduction in infarct size reported in many studies (Abdel-Latif et al., 2007. Arch Intern Med.167:989-997; Bolli et al., 2011. Lancet.378:1847-1857; Donndorf et al., 2011. J Thorac Cardiovasc Surg.142:911-920; Makkar et al., 2012. Lancet.379:895-904; Traverse et al., 2012. JAMA.308:2380-2389). At the same time, the heterogeneity in calcium handling and force generation properties observed between stem cell-derived and primary myocytes result in force transmission impairment and pro-arrhythmic events that ultimately limit the therapeutic benefit (Makkar et al., 2012. Lancet.379:895-904; Traverse et al., 2012. JAMA.308:2380-2389). Equivalents

In describing embodiments of the invention, specific terminology is used for the sake of clarity. For purposes of description, each specific term is intended to at least include all technical and functional equivalents that operate in a similar manner to accomplish a similar purpose.

Additionally, in some instances where a particular embodiment of the invention includes a plurality of system elements or method steps, those elements or steps may be replaced with a single element or step; likewise, a single element or step may be replaced with a plurality of elements or steps that serve the same purpose. Further, where parameters for various properties are specified herein for embodiments of the invention, those parameters can be adjusted up or down by 1/20th, 1/10th, 1/5th, 1/3rd, ½, etc., or by rounded-off approximations thereof, unless otherwise specified. Moreover, while this invention has been shown and described with references to particular embodiments thereof, those skilled in the art will understand that various substitutions and alterations in form and details may be made therein without departing from the scope of the invention; further still, other aspects, functions and advantages are also within the scope of the invention. The contents of all references, including patents and patent applications, cited throughout this application are hereby incorporated by reference in their entirety. The appropriate components and methods of those references may be selected for the invention and embodiments thereof. Still further, the components and methods identified in the Background section are integral to this disclosure and can be used in conjunction with or substituted for components and methods described elsewhere in the disclosure within the scope of the invention.

Claims

We claim:

1. An in vitro method for determining the ability of a stem cell to functionally engraft with a myocyte, comprising

providing a cell co-culture, said cell co-culture comprising

a base layer;

a soft substrate on the base layer;

a biopolymer patterned on the soft substrate;

a single myocyte; and

a single stem cell;

culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface;

determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, and

identifying the stem cell as suitable for functionally engrafting with the myocyte if the contractile force of the stem cell is about the same as the contractile force of the myocyte,

thereby determining the ability of the stem cell to functionally engraft with the myocyte.

2. An in vitro method for identifying a compound useful for functionally engrafting a stem cell and a myocyte, comprising

providing a cell co-culture, said cell co-culture comprising

a base layer;

a soft substrate on the base layer;

a biopolymer patterned on the soft substrate;

a single myocyte; and

a single stem cell;

culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface; contacting the co-culture with a test compound; and

determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for functionally engrafting the myocyte and the stem cell, thereby identifying a compound useful for functionally engrafting the stem cell and the myocyte.

3. An in vitro method for identifying a compound useful for treating a subject having regenerative muscle cell therapy, comprising

providing a cell co-culture, said cell co-culture comprising

a base layer;

a soft substrate on the base layer;

a biopolymer patterned on the soft substrate;

a single myocyte; and

a single stem cell;

culturing the single myocyte and the single stem cell such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface;

contacting the co-culture with a test compound; and

determining a contractile force at the medial cell-to-cell interface of the myocyte and the stem cell, wherein if the contractile force of the stem cell is about the same as the contractile force of the myocyte in the presence of the test compound as compared to the absence of the test compound then the compound is useful for treating a subject having regenerative muscle cell therapy,

thereby identifying a compound useful for treating a subject having regenerative muscle cell therapy.

4. The method of any one of claims 1-3, wherein the stem cell is selected from the group consisting of a primary embryonic stem cell, a stem cell from an embryonic stem cell line, a primary fetal stem cell, a stem cell from a fetal stem cell line, a primary adult stem cell, a stem cell from an adult stem cell line, a stem cell de-differentiated from an adult cell, and an induced pluripotent stem cell (iPS).

5. The method of any one of claims 1-3, wherein the myocyte is selected from the group consisting of a cardiomyocyte, a vascular smooth muscle cell, a smooth muscle cell, a skeletal striated muscle cell, and an airway smooth muscle cell.

6. The method of any one of claims 1-3, wherein the soft substrate has a Young’s modulus of about 1 to about 100,000 pascal (Pa).

7. The method of any one of claims 1-3, wherein the soft substrate has a Young’s modulus of about 1 to about 25,000 pascal (Pa).

8. The method of any one of claims 1-3, wherein the soft substrate is selected from the group consisting of a polyacrylamide gel, a poly(N-isopropylacrylamide), a pHEMA, a collagen, a fibrin, a gelatin, an alginate, and a dextran.

9. The method of any one of claims 1-3, wherein the base layer is selected from the group consisting of a silicon wafer, a glass cover slip, a multi-well plate, a tissue culture plate, and a chamber of a microfluidic device.

10. The method of any one of claims 1-3, wherein the biopolymer is selected from the group consisting of a protein, a carbohydrate, a lipid, and a nucleic acid, or combinations thereof.

11. The method of claim 9, wherein the biopolymer is selected from the group consisting of a fibronectin, a laminin, a collagen, a gelatin, a fibrin, a vitronectin, and a fibrinogen, and combinations thereof.

12. The method of claim 10, wherein the biopolymer is fibronectin.

13. The method of any one of claims 1-3, wherein the soft substrate is functionalized with a bifunctional or a multifunctional linker molecule.

14. The method of claim 13, wherein the bifunctional or multifunctional linker is N- sulfosuccinimidyl 6-hexanoate (sulfo-SANPAH).

15. The method of any one of claims 1-3, wherein the soft substrate is functionalized with a repellent.

16. The method of claim 15, wherein the repellent is agarose, hyaluronic acid, or alginate.

17. The method of any one of claims 1-3, wherein the soft substrate further comprises a fluorescent bead.

18. An in vitro model of myocyte cell therapy, comprising

a cell co-culture, said cell co-culture comprising

a base layer;

a soft substrate on the base layer;

a biopolymer patterned on the soft substrate;

a single myocyte ; and

a single stem cell, wherein the single myocyte and the single stem cell are cultured such that they align substantially anisotropically along their length to the patterned biopolymer and substantially structurally integrate at the medial cell to cell interface.