Tensile properties of vascular smooth muscle cells: Bridging vascular and cellular biomechanics

Journal of Biomechanics 45 (2012) 745–755

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Review

Tensile properties of vascular smooth muscle cells: Bridging vascular and cellular biomechanics Takeo Matsumoto n, Kazuaki Nagayama Biomechanics Laboratory, Department of Mechanical Engineering, Nagoya Institute of Technology, Gokiso-cho, Showa-ku, Nagoya 466-8555, Japan

a r t i c l e i n f o

abstract

Article history: Accepted 30 September 2011

Vascular walls change their dimensions and mechanical properties adaptively in response to blood pressure. Because these responses are driven by the smooth muscle cells (SMCs) in the media, a detailed understanding of the mechanical environment of the SMCs should reveal the mechanism of the adaptation. As the mechanical properties of the media are highly heterogeneous at the microscopic level, the mechanical properties of the cells should be measured directly. The tensile properties of SMCs are, thus, important to reveal the microscopic mechanical environment in vascular tissues; their tensile properties have a close correlation with the distribution and arrangement of elements of the cytoskeletal networks, such as stress fibers and microtubules. In this review, we first introduce the experimental techniques used for tensile testing and discuss the various factors affecting the tensile properties of vascular SMCs. Cytoskeletal networks are particularly important for the mechanical properties of a cell and its mechanism of mechanotransduction; thus, the mechanical properties of cytoskeletal filaments and their effects on whole-cell mechanical properties are discussed with special attention to the balance of intracellular forces among the intracellular components that determines the force applied to each element of the cytoskeletal filaments, which is the key to revealing the mechanotransduction events regulating mechanical adaptation. Lastly, we suggest future directions to connect tissue and cell mechanics and to elucidate the mechanism of mechanical adaptation, one of the key issues of cardiovascular solid biomechanics. & 2011 Elsevier Ltd. All rights reserved.

Keywords: Cellular biomechanics Mechanical properties Cytoskeletal networks Stress fibers

Contents 1. 2.

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4.

n

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 746 Experimental techniques for a tensile test of VSMCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 2.1. Preparation of cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 2.2. Tensile tester . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 2.3. Data analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 Tensile properties of VSMCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 748 3.1. Basic properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 748 3.1.1. Effects of strain rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 748 3.1.2. Effects of differences in the cell detachment method. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 748 3.1.3. Effects of preconditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 748 3.2. Viscoelastic properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 750 3.3. Tensile properties of contracted cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 750 3.4. Effects of phenotype transformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 750 3.5. Anisotropy of a cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 751 Effects of cytoskeletal filaments on the mechanical properties of VSMCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 751 4.1. Mechanical properties of cytoskeletal filaments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 751 4.2. Contribution of cytoskeletal structures to tensile properties of VSMCs. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 751 4.3. Contributions of cytoskeletal structures to intracellular force balance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 752

Correspondence to: Nagoya Institute of Technology, Omohi College, Gokiso-cho, Showa-ku, Nagoya 466-8555, Japan. Tel./fax: þ81 52 735 5049. E-mail address: [email protected] (T. Matsumoto).

0021-9290/$ - see front matter & 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.jbiomech.2011.11.014

746

5.

T. Matsumoto, K. Nagayama / Journal of Biomechanics 45 (2012) 745–755

Future directions and conclusion Conflict of interest statement . . . Acknowledgment . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . .

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1. Introduction Smooth muscle cells (SMCs) are a type of muscle cell found within the walls of organs, such as intestines, bronchi, and blood vessels. They are called ‘smooth’ because they lack the striations seen in the striated muscle. Compared with skeletal muscle cells, SMCs are small in size, slow in response, and highly resistant to fatigue. In vascular walls, vascular smooth muscle cells (VSMCs) reside mostly in the media and align circumferentially. They have a spindle shape and a typical size of  100 mm in length and 5 mm in diameter in the wall, with actin filaments aligning in their axial direction with a slightly oblique arrangement (Fig. 1A). VSMCs play various roles in the vascular wall: their contraction and relaxation change the vascular diameter and control local blood flow at the arteriolar level. It has been indicated that the characteristic impedance of the artery is maintained at minimum levels with the contraction of VSMCs (Cox, 1975, 1976) and that their contraction and relaxation might control intramural stress distribution (Matsumoto et al., 1996). When exposed to hypertension, the cells become hypertrophied to maintain a circumferential stress under physiological conditions (Berry and Greenwald, 1976; Vaishnav et al., 1990; Matsumoto and Hayashi, 1994, 1996). In the course of atherogenesis, the VSMCs proliferate, migrate into the subendothelial space, secrete extracellular matrix (ECM), take up lipids, and become foam cells (Raines and Ferri, 2005). VSMCs can be isolated from vascular tissues by enzymatic digestion (Chamley et al., 1977) or an explant method (Ross, 1971). When cultured, the cells change their phenotype from contractile to synthetic (Campbell and Campbell, 1995). Contractile cells are the cells found in the normal arterial wall, and they have abundant myofilaments and few organelles (Fig. 1A); they contract and relax in response to mechanical and biochemical

Fig. 1. Fluorescent images of the stress fibers of rat aortic SMCs. A freshly isolated SMC (A), a freshly isolated SMC with contraction induced by serotonin (B), and a cultured SMC isolated from substrate (C). Note that the obliquely aligned fibers of freshly isolated VSMC (A, arrow) changed into the aggregated fibers with some buckled structures (B, arrowheads). The fibers in cultured VSMCs isolated from substrate showed entangled-basket-like structures (C).

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. 752 . 753 . 753 . 753

signals (Fig. 1B), have little synthetic ability, and seldom proliferate. In contrast, synthetic cells are found in development and repair processes, such as in atherosclerotic lesions, and their cytoplasm contains few filament bundles but large amounts of organelles, including rough endoplasmic reticulum (RER) and Golgi. Their function is almost exclusively related to protein synthesis; they also proliferate, but hardly contract in response to contractile agonists. As indicated by the wall thickening that occurs in hypertension, it is highly probable that the mechanical environment has profound effects on the activities of SMCs. The active contraction following passive extension has long been known as the Bayliss effect (Bayliss, 1902). Cyclic deformation of the substrate has various effects on cultured SMCs (Williams, 1998), such as cell alignment, increased protein synthesis and proliferation, and stiffening (Smith et al., 2003; Na et al., 2008). These responses are accompanied by rapid changes at the subcellular level: the remodeling of integrins occurs within a minute in response to cyclic stretch (Cunningham et al., 2002) and is accompanied by a temporal increase in cell stiffness and focal adhesion area (Na et al., 2008). Rapid mechanical responses are also observed at a tissue level; the VSMCs within the wall of arterioles shorten during the initial vasoconstriction caused by norepinephrine (NE) and then elongate to restore their original length during prolonged exposure to NE within 4 h (Martinez-Lemus et al., 2004). To study the relationship between such mechanical stimulations and these dynamic events occurring in the artery wall, the intramural mechanical environment at the cellular level should be established. Estimations of the cell mechanical environment have been studied in fibroblasts cultured in gels (Brown et al., 1998; Nekouzadeh et al., 2008). In the case of gels that have a relatively homogeneous structure and an elastic modulus almost comparable to that of cells, reliable estimation may be obtained for the mechanical environment of cells by comparing the tensile properties of the gels with and without cells. However, arterial walls have a highly heterogeneous structure at the microscopic level, which complicates a mechanical analysis. For example, the media of the elastic arteries, such as aortas, have a layered structure called a lamellar unit, which is a pair of elastic lamina composed of elastin and a smooth muscle-rich layer mainly composed of VSMCs and collagen (Wolinsky and Glagov, 1967). The elastic modulus of elastin is  0.6 MPa (Fung, 1981), whereas that of smooth muscle is of the order of 10 kPa in the relaxed state (Whitemore, 1968), and that of collagen is 1 GPa (Fung, 1981). Such heterogeneity may cause a heterogeneous distribution of intramural stress in the loaded wall (Matsumoto et al., 2008) and a complicated distribution of residual stress (Matsumoto et al., 2004). Thus, stress and strain distributions in the artery wall appear to be highly complicated at the microscopic level. To elucidate the mechanical environment of VSMCs precisely, we, therefore, need to measure the mechanical properties of VSMCs isolated from artery walls, while considering their in situ strain field in the wall. Because VSMCs are cyclically stretched by  10% with a 25–50% mean strain (Matsumoto et al., 2008) in the cardiac cycle, their mechanical properties should be determined during a wide range of deformations. The mechanical properties of cells have been extensively studied by micropipette aspiration (Hochmuth, 2000), magnetic particle twisting (Wang and Ingber, 1994; Smith et al., 2003), and

T. Matsumoto, K. Nagayama / Journal of Biomechanics 45 (2012) 745–755

nanoindentation with atomic force microscopy (AFM) (Hoh and Schoenenberger, 1994; Sato et al., 2000; Ludwig et al., 2008) mainly using cultured cells. However, these studies only reported on the mechanical properties of local regions of cells under small deformation and did not provide sufficient information concerning the magnitude of the stress applied at the whole-cell level under a physiological strain range. We, therefore, need to obtain quantitative data on the tensile properties of VSMCs that have been freshly isolated from the vessel walls; these data are expected to be important from the viewpoint of tissue biomechanics. Tensile testing is also important for cultured cells from the viewpoint of cell biomechanics. Most of the fundamental questions of cell biomechanics include how a macroscopic deformation of a cell is transmitted to its organelles, how their deformation mediates its mechanotransduction, and how its mechanical properties are affected by the organelles. Membranous organelles, such as RER and Golgi, may have minor effects on the mechanical properties of the cells, but their deformation might have significant effects on signal transduction by changing the distance, i.e., transmission efficiency of second messengers, between the organelles and the molecular machines associated with them. Tensile testing that can impose a large deformation is very useful for this purpose. In such experiments, we may need to visualize a particular organelle to know its deformation, and to destroy it to determine its effects on whole-cell mechanics. These techniques can be applied more easily and efficiently in cultured cells. In this review, we will first introduce the experimental techniques used for the tensile testing of VSMCs, both freshly isolated and cultured, and then discuss various factors affecting the tensile properties of VSMCs. As cytoskeletal networks are particularly important for the mechanical properties of a cell and its mechanotransduction, the mechanical properties of cytoskeletal filaments and their effects on whole-cell mechanical properties are discussed paying attention to the intracellular force balance. Lastly, we suggest future directions to connect tissue and cell mechanics and to elucidate the mechanism of mechanical adaptation, one of the key issues of cardiovascular solid biomechanics.

2. Experimental techniques for a tensile test of VSMCs 2.1. Preparation of cells Two types of cells are used depending on the purpose of the experiment. One comprises VSMCs freshly isolated from vascular tissues by enzymatic digestion, and the other comprises cultured VSMCs. Freshly isolated cells retain their spindle shape, and their typical dimensions are 20–100 mm in length and 5–10 mm in diameter. When measuring the tensile properties of cultured cells, we need to pay attention to the cell detachment method because their shape and internal structure is completely different with the different methods used to detach the cells from the substrate (Fig. 2; see Section 3.1.2). 2.2. Tensile tester Cells are usually gripped with a pair of microtools under a microscope and stretched horizontally by moving one of the microtools with a micromanipulator (Fig. 2). Table 1 summarizes the gripping methods along with their advantages and disadvantages. If the cells are long enough (several 100 mm), such as those obtained from the stomach, they can be gripped by knotting each end around a micropipette tip. For shorter cells, they are gripped

747

Fig. 2. Schematic of the cell gripping and tensile testing of a trypsinized cell (A) and a cell maintaining its shape on the substrate (B).

by aspirating them at both ends into micropipettes, or by gently pressing the micropipettes, microrods, or microplates onto their lateral surface. In the latter method, the cells are stretched at a portion of their surface, which might cause shear deformation of the cells. However, the stiffness obtained in the above methods has been reported to be similar (Nagayama et al., 2006). The lateral surface-gripping method has been widely used for its technical easiness but also for its versatility, for example, the cells can be stretched in the lateral direction (Section 3.5) and adherent cells can be stretched while maintaining their shape on the substrate (Section 3.1.2). The surface of the microtools are often coated with various adhesives, including a cellular adhesive (Miyazaki et al., 2000) and a urethane adhesive (Shue and Brozovich, 1999), to improve the adhesion; uncoated carbon fibers are also used for their high affinity to cardiomyocytes (Sugiura et al., 2006). Tension T applied to a cell is measured by the deformation x of a microcantilever whose spring constant k has been calibrated (Matsumoto et al., 2000) as T¼kx. However, a calibration is necessary every time the pipette is changed. To overcome this problem, tensile testers with handmade load cells have also been proposed (Glerum and Van Mastrigt, 1990; Miyazaki et al., 2000). SMCs are also highly viscous, and a stress relaxation test is useful to measure their viscoelasticity, in which the cell length needs to be kept precisely constant. For this purpose, tensile testers with visual feedback control have been proposed (Iribe et al., 2007; Nagayama et al., 2007b). 2.3. Data analysis A tension–elongation curve of the specimen cell is immediately obtained by plotting the tension T against its elongation DL calculated as the increment of L, the distance between the pipettes (Fig. 2). The linearized strain e is obtained by normalizing the elongation DL with the initial distance between the two pipettes L0. Cell mechanical properties have been evaluated using the slope of the tension–elongation or tension–strain curves, namely, stiffness S. A stress–strain relationship can also be obtained, but it is not easy because 3D finite element analysis considering a large deformation is required for the rigorous analysis and 3-D shapes of the cell are necessary to build the finite element model. As a preliminary analysis, nominal stress– strain relationships have been obtained. The nominal stress s is obtained by dividing tension T with the initial cross-sectional area A0 before the stretch. The initial cross-sectional area has been

748

T. Matsumoto, K. Nagayama / Journal of Biomechanics 45 (2012) 745–755

Table 1 Summary of the cell gripping method for the tensile test for SMCs. Microtool

Gripping method

Pipette

Image

Easy Without grippinga shearb

Lateral In situ Other stretchc stretchd advantage(s)

Other Ref. disadvantage(s)

Knotting

N

Y

N

N

Limited only for very long cells

Pipette

Aspiration

NN

Y

Y/N

N

Pipette or rod

Adhesion

YY

N

Y

Y

Plate

Adhesion

Y

Y

Y

N

Micropillar Adhesion array substrate

–

N

Y

Y

Strong grip

Warshaw and Fay (1983); Warshaw et al. (1986); Glerum and Van Mastrigt (1990b) Matsumoto et al. (2000); Miyazaki et al. (2000)

Shue and Brozovich (1999); Smith et al. (2000); Nagayama et al. (2006); Sugiura et al. (2006)

No need to grip, just stretch the substate

Weak gripping

Thoumine and Ott (1997)

Unsuitable for large deformation

Nagayama et al. (2011) (See Chapter 5 for details)

a

Is cell gripping easy? (Yes: YY 4Y4N 4NN: No). Is it possible to stretch cells without shear deformation? (Y, possible; N, impossible). c Is it possible to stretch cells in the direction perpendicular to their longitudinal axis? (i.q.). d Is it possible to stretch cells while maintaining their shape on a culture dish? (i.q.). b

obtained in various ways. For example, in the case of cells with a round cross-section, A0 is obtained as A0 ¼ p(D0/2)2, where D0 is the nominal diameter of the cell obtained by dividing the initial tracing area of the cell S0 with the initial distance between the two pipettes L0. The mechanical properties of the cells are then evaluated with the normalized stiffness, that is, the slope of the nominal stress–strain curves. The stiffnesses of the VSMCs obtained in such analyses under various conditions are summarized in Table 2.

3. Tensile properties of VSMCs 3.1. Basic properties 3.1.1. Effects of strain rate Because SMCs show viscoelasticity, their tensile properties are sensitive to the strain rate. As an example, the normalized stiffness (kPa) of cultured bovine aortic SMCs increased linearly with the strain rate (%/s) with a proportionality factor of 0.495 (Matsumoto et al., 2005). Attention should be given to the strain rate when comparing the data obtained in different experiments. In our studies, we have been stretching cells at a rate of 1 mm for 5 s, which corresponds to o4%/s depending on the initial length L and the spring constant k, where the strain rate effect is minor. 3.1.2. Effects of differences in the cell detachment method Cultured cells are usually detached from culture dishes by trypsinization for tensile tests. Trypsinized cells become round following detachment and lose their shape and cytoskeletal integrity on the substrate (Figs. 1C and 2A), suggesting that significant changes in their mechanical properties may occur. To avoid such changes, we developed a new method, a quasi-in situ

tensile test, to stretch the cells, while maintaining their shape on the substrate (Nagayama and Matsumoto, 2008). The VSMCs on the substrate are treated with a Ca2 þ –Mg2 þ -free Hank’s balanced salt solution to inhibit cell contraction and morphological changes with the mechanical stimuli and the cell adhesiveness to the substrate. A pair of micropipettes coated with an adhesive is then pressed onto the surface of the cell body at both end regions (Fig. 2B), and the target cell is detached by carefully lifting up the pipettes while maintaining the cell’s in situ shape on the substrate. The stiffness S obtained in the quasi-in situ tensile test (Table 2 #14 and 15) was higher than that obtained for trypsinized spherical VSMCs (Table 2 #1 and 3). Furthermore, the normalized stiffness was much higher in the cells with an in situ cell shape (Table 2 #14 and 15) than in the trypsinized cells (Table 2 #1 and 3). The actin filaments in trypsinized cells look aggregated and entangled (Fig. 1C), and they align in the major axis direction of the cell on the substrate. Such a difference may cause a softening of trypsinized cells.

3.1.3. Effects of preconditioning Although this need has long been pointed out, specimen cells were not preconditioned in the tensile test for years due to technical difficulties (Thoumine and Ott, 1997; Matsumoto et al., 2000; Miyazaki et al., 2000; Nagayama et al., 2006). We have recently succeeded in observing a change in the tensile properties during cyclic stretching (Nagayama and Matsumoto, 2008). Typical examples of tension–strain curves are shown in Fig. 3A. The curves in the range of e 45% of the net strain were relatively linear. The hysteresis area of the loading/unloading process was relatively large in the first cycle but decreased significantly and became stable in the second and subsequent cycles: the VSMC was mechanically preconditioned at the first

Table 2 Summary of the stiffness of VSMCs obtained in tensile tests under various conditions. #

Aminal Conditions

n

Strain rate (%/s)

Fresh or cultured

Shape

Stretch direction

Preconditioned?

Cell treatment

Stiffness (N/m)

Stiffness (nN/%)

Normalized stiffness (kPa)

Overall

Initial (e o 0.2) Overall

Initial (e o 0.2)

Overall

0.027 70.010 0.036 70.021 0.092 70.071 0.055 70.033 0.097 0.027 0.17 70.032 0.88 70.45 0.065 70.037

5.67 3.3 15.1 721.5 16.2 715.7 8.47 7.4 – – – 16.2 79.4

2.75 7 0.92 12.6 7 10.8 3.4 73.0 10.4 79.1 – – – 14.8 7 9.6

1.51 7 0.49 9.3 77.8 2.6 72.9 11.0 75.4 – – – 13.6 7 10.8

1 2 3 4 5 6 7 8

Bovine Rat Rat Rat Rabbit Rabbit Rabbit Rat

0.2–3.4 0.2–2.6 0.27–0.82 0.21–0.63  30  30  30 0.18–0.76

Cultured Fresh Cultured Fresh Cultured (Fresh)a (Fresh)a Fresh

Round Fusiform Round Fusiform Round Round Round Fusiform

NA Major NA Major NA Major Major Major

No No No No No No No No

None None None None None None Norepi. None

6 8 21 8 6 6 6 5

0.048 þ0.020 0.062 þ0.058 0.106 70.064 0.049 70.041 – – – 0.070 7 0.036

9

Rat

0.13–0.95

Fresh

Fusiform Minor

No

None

5

0.097 70.056 0.068 70.081 (72.6 751.1)c

(51.37 71.1)c

10 Rat

0.16–0.55

Fresh

Fusiform Major

No

Serotonin

4

0.595 7 0.234 0.611 70312

96.77 58.6

11 Rat

0.14–0.58

Fresh

Fusiform Minor

No

Serotonin

4

0.187 7 0.092 0.131 70.109 (151.3 776.7)c (103.37 67.4)c

12 Rabbit 13 Rabbit

0.08–0.71 0.04–0.21

Fresh Fresh

Fusiform Major Fusiform Major

No No

None Serotonin

6 8

14 Rat

0.09–0.15

Cultured

In situ

Major

No

15 Rat

0.04–0.10

Cultured

In situ

Major

16 Rat

0.04–0.13

Cultured

In situ

17 Rat

0.07–0.16

Cultured

18 Rat

0.06–0.12

19 NT Rat 0.4–1.8 20 HT Rat 0.6–2.3

93.1 752.3

3.17 1.4 8.17 8.4 13.17 10.3 9.67 4.4 ( 18)b ( 34)b ( 170)b 14.97 9.6

None

0.055 70.036 0.0507 0.030 13.4 75.6 0.659 7 0.564 0.589 70.594 98.3 753.0 (e o 5%) 10 – – 8.17 3.1

12.37 5.9 87.17 58.9 (5%o e) 14.97 4.4

Yes

None

10 –

–

13.07 3.4

23.37 5.4

Major

Yes

Serotonin

6

–

–

27.3 73.1

39.67 5.1

In situ

Major

Yes

Cyto-D

9

–

–

5.17 2.6

8.67 2.9

Cultured

In situ

Major

Yes

Colchicine

9

–

–

13.07 2.9

18.17 4.3

Fresh Fresh

Fusiform Major Fusiform Major

No No

None None

5 6

12.3 79.6 6.27 3.0

10.47 4.9 4.97 3.0

Matsumoto et al. (2000) Matsumoto et al. (2000) Nagayama et al. (2006) Nagayama et al. (2006) Miyazaki et al. (2002) Miyazaki et al. (2002) Hayashi (2006) Nagayama and Matsumoto (2004) 2.8 72.4 1.8 73.0 Nagayama and Matsumoto (2004) 88.1 7 26.6 92.4 7 30.1 Nagayama and Matsumoto (2004) 59.6 7 18.8 38.3 7 25.8 Nagayama and Matsumoto (2004) 15.5 7 6.9 14.4 7 7.5 Original (Fig. 4A) 156.4 7 105.9 121.4 7 81.3 Original (Fig. 4B) (e o 5%) (5%o e) (11)d (21)d Nagayama and Matsumoto (2008) (18)d (32)d Nagayama and Matsumoto (2008) – – Nagayama and Matsumoto (2010) – – Nagayama and Matsumoto (2008) – – Nagayama and Matsumoto (2008) 10.2 74.9 8.1 76.2 Matsumoto et al. (2011) 4.3 71.4 3.4 71.6 Matsumoto et al. (2011)

T. Matsumoto, K. Nagayama / Journal of Biomechanics 45 (2012) 745–755

Initial (e o 0.2)

Refa

NA, not applicable; Norepi, norepinephrine; Cyto-D, cytochalasin D; NT, normotensive; HT, hypertensive. a

Cultured for 3 days after isolation by enzyme. Rough estimation assuming initial cell length to be 20 mm. Values are high because of a larger cross-sectional area and shorter cell length due to stretch along the short axis. d Estimated with a cross-sectional area of SMCs on a substrate perpendicular to the stretch direction (727 17 mm2, n¼23). b c

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Fig. 4. Tensile properties of smooth muscle cells freshly isolated from rabbit thoracic aortas. Nominal tension–strain curves of untreated cells (A) and cells exposed to 10  5 M serotonin (B).

cycle of the loading/unloading process, and the force–strain curve became stable following the second cycle. The slope of the loading curves increased, i.e., the cell stiffened, by 54% between the first and second stretch and did not change significantly afterwards (Table 2 #14 vs. #15).

3.2. Viscoelastic properties Local viscoelastic properties have been investigated extensively for airway SMCs with magnetic bead twisting (Stamenovic et al., 2002; Stamenovic, 2008) and AFM (Smith et al., 2005), and sophisticated models, such as a power-law structural dumping model, have been proposed. In contrast, there is a paucity of viscoelastic data for whole cells. To the authors’ knowledge, only stress relaxation tests have been conducted for VSMCs (Nagayama et al., 2007a), in which a reduced relaxation function (Fung, 1981) that can be fitted as Gðt, sÞ ¼ 0:71ðet=6 þet=65 þ et=530 Þ þ 0:29 was obtained for cultured rat aortic SMCs following rapid stretching with a strain rate of 10%/s. Normalized stiffness measured in the rapid stretch phase was 12.772.2 (kPa, mean7SEM, n¼11) and was almost 4 times the stiffness obtained in the quasi-static tensile test (Table 2 #3). The strain rate in the artery wall during pulsation is  10%/s. Considering that elastin is almost an elastic material (Fung, 1981), VSMCs may experience a much higher stress than expected from static analyses during the pulsation of the wall.

3.3. Tensile properties of contracted cells Fig. 4 shows examples of the tensile properties of relaxed and contracted VSMCs isolated from rabbit thoracic aortas. A maximal contraction was induced with 10  5 M serotonin, and the initial normalized stiffness increased 10-fold in response to contraction (Table 2 #12 vs. #13). Similar results were obtained for VSMCs isolated from rat thoracic aortas (Table 2 #8 vs. #10). These results indicate that the tension borne by the VSMCs in the aortic wall increases several times upon their maximal contraction. In contrast, the artery diameter decreases upon smooth muscle contraction, indicating that the circumferential strain applied to other components in the vessel wall, such as collagen and elastin, may decrease. Taken together, the proportion of the stress borne by VSMCs may increase significantly upon smooth muscle contraction. 3.4. Effects of phenotype transformation Significant softening occurs when contractile VSMCs change to a synthetic state (Nagayama et al., 2006). For SMCs obtained from rat thoracic aortas, their normalized stiffness decreases to onethird or one-fourth upon phenotype transformation (Table 2 #3 vs. #4). Contractile cells have abundant actin filaments and few organelles, whereas synthetic cells contain few filament bundles but large amounts of organelles. As shown in Section 4,

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cytoskeletal filaments are much stiffer than the cytoplasm, whereas other organelles are supposed to be highly deformable because they have a membranous structure without a lining of cytoskeletal networks. Such differences in structure may explain the softening observed for the synthetic cells. Similar results have been obtained for SMCs derived from rabbit thoracic aortas (Miyazaki et al., 2002); however, the decrease in the stiffness upon phenotype transformation was less than 50% (Table 2 #5 vs. #6). The cells used in this study were isolated and cultured for 3 days. Although these cells showed a contractile response and expression of the smooth muscle myosin heavy chain, these cells might have been partly synthetic. Recently, we found that the normalized stiffness of rat aortic SMCs decreased by 60% (Table 2 #19 vs. #20) in response to 16-week hypertension (Matsumoto et al., 2011). Studies have reported that hypertension induces change similar to phenotypic transformation from contractile to synthetic (Guyton et al., 1990). The softening of hypertensive SMCs might be caused by the phenotypic transformation. 3.5. Anisotropy of a cell Contractile VSMCs are spindle-shaped and their intracellular contractile apparatus, such as actin and myosin filaments, run mostly parallel to their major axis. As will be shown in Section 4, actin filaments dictate cell stiffness. Thus, VSMCs may be stiffer in their major axis. To test this hypothesis, the tensile properties of VSMCs freshly isolated from rat aortas were measured in their major and minor axes (Nagayama and Matsumoto, 2004). Their normalized stiffness was significantly higher in the major axis than the minor axis (Table 2 #8–11). We also found that the stiffness increased significantly and the difference between the major and minor axes disappeared in contracted cells. Anisotropic alignment of actin filaments disappears in contracted cells (Fig. 1B). This may explain the isotropy of the contracted cells.

4. Effects of cytoskeletal filaments on the mechanical properties of VSMCs VSMCs contain three main kinds of cytoskeletal filaments: actin stress fibers (SFs), microtubules (MTs), and intermediate filaments (IFs). These cytoskeletal filaments play dominant roles in various cellular events including cell migration (Haga et al., 2000), proliferation (Ingber, 1990), differentiation (Collinsworth et al., 2002), and apoptosis (Chen et al., 1997), and they determine the mechanical properties and shape stability of cells. They are also believed to play pivotal roles in the process of intracellular mechanotransduction (Wang et al., 1993; Davies, 1995). To study the mechanism of mechanotransduction, it is crucial to determine the mechanical properties of cytoskeletal filaments and their contribution to the mechanical properties of cells. In this section, we discuss the basic structural features and mechanical properties of cytoskeletal filaments and their roles in the tensile properties of VSMCs. 4.1. Mechanical properties of cytoskeletal filaments SFs are composed of bundles of actin filaments (AFs). These bundles are held together by the actin-crosslinking protein a-actinin, and they contain non-muscle myosin and tropomyosin (Pellegrin and Mellor, 2007). SFs generate intracellular tension through actomyosin activation, which is recognized as the fundamental mechanism for tension development in adherent contractile cells, including VSMCs. These contractile forces are transmitted to ECM via focal adhesions, and the contractile properties of single SFs have been investigated quantitatively (Katoh et al., 1998). These

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investigators isolated SFs from endothelial cells and fibroblasts with a low-ionic-strength extraction solution and showed that SFs shortened  20% of the initial length in the presence of Mg-ATP. Deguchi et al. (2006) isolated SFs from cultured bovine VSMCs using a similar technique and measured the tensile properties of single SFs by in vitro manipulation with carbon fibers. They then estimated Young’s modulus of the isolated SFs to be 1.45 MPa, which was approximately three orders of magnitude lower than that of single AFs (1.8–2.6 GPa) (Kishino and Yanagida, 1988; Kojima et al., 1994; Liu and Pollack, 2002). The stress–strain curve of the single AFs was highly linear, whereas that of the SFs showed a remarkable nonlinear behavior or strain-induced hardening. The breaking force of single AFs was reported to be 600 pN (Tsuda et al., 1996), whereas that of the single SFs increased more than 600 times (  380 nN) (Deguchi et al., 2006). The tension applied to the focal adhesions at the cell periphery increased from several 10 nN to as much as  100 nN upon contraction of the VSMCs (Nagayama and Matsumoto, 2011). Thus, the SFs are strong enough to bear the maximum contractile force. Each of the SFs also stiffened in response to contraction. This was associated with actomyosin activation and also with the fusion of SFs (Nagayama and Matsumoto, 2010). MTs are hollow biopolymers (Nogales, 2000) whose inner and outer diameters are 14 and 25 nm, respectively. Intracellular MTs are composed of tubulin dimers organized at the centrosome; they are crucial for mitosis (Pickett-Heaps and Northcote, 1966) and for the dynamic positioning of intracellular organelles and the nucleus (Robinson et al., 1995; Tolic´-Nørrelykke, 2008). MTs are the most rigid cytoskeletal filaments, with a bending rigidity 4100 times that of AFs (Gittes et al., 1993), and in certain instances, they attach to the cell membrane via certain capping proteins (Reilein and Nelson, 2005). Thus, microtubules play important roles in the stabilization of cell elongation, including nerve cell extension (Zheng et al., 1993). Young’s modulus of single MTs has been estimated to be 1.2 GPa by either recording thermally induced shape fluctuations (Gittes et al., 1993) or applying a bending force to MTs (Kurachi et al., 1995; Kis et al., 2002), which is the same order of magnitude as that of single AFs. IFs, with a diameter of 10 nm, are sufficiently compliant to generate moderate deformation, and they maintain their resistance to large deformation to provide a structural integrity in cells. The VSMCs contain vimentin as the most prominent IF, whereas desmin, a marker for muscle cells, is also present in small vessels (Wede et al., 2002). The vimentin filaments form a dynamic network and change from curved-fiber networks with random directions to straight-fiber networks along the cell major axis following contractile stimulation (Tang et al., 2005). Young’s modulus of single vimentin filaments has been estimated to be 300–900 MPa using bending experiments with AFM (Guzma´n et al., 2006). 4.2. Contribution of cytoskeletal structures to tensile properties of VSMCs Information concerning the contribution of the cytoskeletal structures to the mechanical properties of cells under large deformation is especially important for the VSMCs. As discussed in previous sections, a single cell tensile test maintaining in situ cell shape and cytoskeletal integrity is a powerful technique to investigate the mechanical properties of the VSMCs, and a comparison of the cell stiffness before and after the disruption of each cytoskeletal network is useful for a study of their mechanical contribution (Nagayama and Matsumoto, 2008, 2010). These tests revealed that the SFs were the most essential component in maintaining the tensile properties of the VSMCs: disruption of the SFs decreased the tensile stiffness of the VSMCs

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by 450% (Table 1 #15 vs. #17). Disruption of the MTs also decreased the tensile stiffness of the VSMCs by  30% in the large strain range (e 45%) but did not have an effect in the small strain range (Table 1 #15 vs. #18). Because intracellular MTs show a wavy morphology under no-load conditions, it is, therefore, conceivable that the MTs are partly under tension and act as a resistance to stretch when the strain of the cell is relatively large. The SFs and MTs have significant effects on the viscoelastic properties of the VSMCs. The hysteresis of the force–strain curve obtained in the loading/unloading experiments, which represents the viscous energy dissipation, increased dramatically following SF disruption (Fig. 3B) and MT augmentation (Nagayama and Matsumoto, 2008). These results may indicate that the SFs and MTs maintain the elastic and viscous properties of the VSMCs, respectively. However, the energy dissipation during cell deformation at a low strain rate might not be due to the passive fluid viscous dissipation (Bursac et al., 2005; Deng et al., 2006) but rather the active reorganization of cytoskeletal networks (Matthews et al., 2006). This type of active reorganization of cytoskeletal networks may have contributed to a part of the viscoelastic responses observed in the mechanical tests. The mechanical contribution of the IFs to the tensile properties of the VSMCs remains to be clarified because, in many cases, IF disruption using acrylamide also disturbed the AF structure in the VSMCs. These responses may be due to the interaction between the IFs and AFs, as observed in epithelial cells and fibroblasts (Green and Goldman, 1986; Green et al., 1987). It has been reported that the vimentin-deficient fibroblasts obtained from gene-disrupted mouse embryos were softer than wild-type cells, except at the lowest applied stress (Wang and Stamenovic´, 2000). This result may indicate that the IFs play important roles in the mechanical properties of the cell during large deformation.

Fig. 5. Intracellular nuclei in the VSMCs were compressed by forces generated by stress fibers; these forces resulted in changes in the 3-D morphology of the nuclei, depending on the intracellular tension exerted by the stress fibers (A). Cell height at the nuclear region decreased from 3.9 mm (B) to 3.3 mm following stress fiber contraction (C), and increased to 10.3 mm following stress fiber disruption (D) (Nagayama and Matsumoto, 2008).

4.3. Contributions of cytoskeletal structures to intracellular force balance A consideration of the intracellular force balance is important for a better understanding of cell mechanics and mechanotransduction. The effects of SFs and MTs on the intracellular force balance have been discussed mainly with a two-dimensional traction force exerted by cells onto a substrate, which was estimated by the deformation of a polyacrylamide gel substrate for airway SMCs (Smith et al., 2000; Stamenovic´ et al., 2002) and the deformation of VSMCs following the noninvasive detachment from a thermoresponsive-gelatin coated substrate (Nagayama and Matsumoto, 2008). These studies demonstrated that the intracellular tension increased with actomyosin activation and decreased with SF disruption, whereas it increased with MT disruption and decreased with MT augmentation. Such reports indicated that the organization of cytoskeletal networks could be expressed by a tensegrity model (Ingber, 1993), in which the SFs are a prestretched elastic component generating tension, and the MTs and ECM oppose the tension. Furthermore, the intracellular MTs have been considered resistant to large-scale compression because of the lateral reinforcement from other elements of the cytoskeletal filaments (Brangwynne et al., 2006). These mechanical contributions of SFs and MTs affect the three-dimensional intracellular force balance in VSMCs and may change the mechanical environment around the nuclei: cell height in the nucleus region decreased and increased following actomyosin activation and SF disruption, respectively (Fig. 5), and decreased and increased following MT disruption and augmentation, respectively (Nagayama and Matsumoto, 2008). It has been suggested that the cytoskeleton has the potential to interact with the nucleus via nuclear membrane proteins such as nesprin and SUN proteins (King et al., 2008), and evidence for a mechanical interaction between the SFs and the nucleus was recently obtained experimentally using a laser

Fig. 6. A mechanical interaction between stress fibers and the nucleus. A single stress fiber running across the top surface of the nucleus was cut by irradiation from a pulsed laser. The inset images indicate the fluorescent images of the nucleus before and after the stress fiber dissection. The stress fibers shortened across the top surface of the nuclei after their dissection. The nuclei moved in the direction of the stress fiber retraction and showed a marked local deformation, indicating that the stress fibers were firmly connected to the nuclear surface.

nanodissection technique (Fig. 6) (Nagayama et al., 2011). The findings indicated that this mechanical interaction may achieve direct force transmission from the SFs to the nucleus and vice versa and may affect the gene expression of VSMCs (Verstraeten and Lammerding, 2008). Vimentin IFs also have the potential to contribute to the intracellular force balance. Vimentin connects to the cytoplasmic dense bodies, which the AFs also connect to, and to desmosomes. It is possible that a special reorientation of vimentin may induce the reorganization of the AFs and strengthen the interaction with the desmosomes, which may regulate tension development in smooth muscle (Tang, 2008; van den Akker et al., 2010). The importance of the IFs for the intracellular force balance of VSMCs needs to be clarified.

5. Future directions and conclusion To elucidate the mechanical adaptation of blood vessel walls, a multiscale mechanical analysis (Lim et al., 2006; Humphrey, 2008) to determine how the deformation applied to tissues is transmitted to cells, to organelles and, ultimately, to molecules is necessary.

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There have been ample data on the mechanical responses at the tissue and cellular levels. Many studies at the subcellular level with recent technologies, such as AFM (Smith et al., 2005) and optical magnetic twisting cytometry (OMTC), have been recently performed. For example, Smith et al. (2003) studied the local stiffness of airway SMCs subjected to cyclic stretch with OMTC and found that the long-term (10–12 days) oscillatory loading caused the stiffness to increase and become anisotropic. Park et al. (2010) visualized the distribution of cytoskeletal prestress in airway SMCs with OMTC and AFM in combination with finite element analysis. These studies at the cellular and subcellular levels are involved with cultured cells of the synthetic phenotype. As shown in Section 3.4, the mechanical properties and structures of synthetic cells are completely different from those of contractile cells. The mechanical environment is also different between the SMCs in culture (2-D) and in tissues (3-D). Furthermore, the composition and stiffness of extracellular matrices are also not similar. When extrapolating the mechanical response of cells in tissues from the data obtained for cells in culture, careful attention should be given to such differences. Thus, the tensile properties and mechanical responses of freshly isolated cells under physiological (large) deformation are crucial to connect vascular biomechanics with cell biomechanics. Knowledge of the intracellular force balance is also an important component in connecting cell biomechanics with molecular biomechanics. To provide more reliable data for multiscale approaches, we still need to overcome many problems. One problem is that freshly isolated VSMCs held between a pair of micropipettes rarely show contraction, even with the softest pipette with a spring constant k of 10 nN/mm, yet, they contract by 20–30% in response to contractile agonists when they have no constraints. A reason for this might be the difference in the load transmission points between the in situ conditions and those of the tensile test conditions. The contractile force developed by cells in situ is transmitted to the extracellular matrix through focal adhesions distributed on the cell surface, whereas it is transmitted only through the area adhering to the micropipettes for cells during the tensile test. This may significantly reduce the force transmission efficiency, and the cells might not show noticeable contraction even with slight constraints. Recently, we found that the VSMCs cultured on an elastic micropillar array substrate actively contracted in response to stretching of the substrate (Nagayama et al., 2011), although the micropillars were stiffer (k¼26 nN/mm) than the micropipettes. A new method to hold a cell on its entire surface is necessary. Another problem is that we do not know the tone of the VSMCs in the tissue. As shown in Section 3.3, the stiffness of the SMCs changes drastically in response to contraction. Thus, we need to estimate the contractile state of the VSMCs in the tissue. As intramural VSMCs change their dimensions within an hour in response to changes in their mechanical environment (MartinezLemus et al., 2004), attention should be given to the changes of the cells that might occur upon isolation. We also need to accumulate knowledge concerning the dynamic deformation of the cytoskeletal networks and organelles during stretching to determine the intercellular force balance; in particular how the deformation of a cell is transmitted to a deformation of the nucleus and how this might have a crucial effect on gene transcription (Wang et al., 2009) should be determined. We anticipate the elucidation of the mechanism of the mechanical adaptation of vascular tissues through the use of these approaches, and one of the key techniques is the tensile test.

Conflict of interest statement The authors declare they have no conflict of interest in their manuscript.

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