This work was supported by the following funding sources: R01-AR063649, AG-020591, F31-AR035931.

<ol>
	<li>Mendias, C. L., Kayupov, E., Bradley, J. R., Brooks, S. V., Claflin, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decreased+specific+force+and+power+production+of+muscle+fibers+from+myostatin-deficient+mice+are+associated+with+a+suppression+of+protein+degradation.">Decreased specific force and power production of muscle fibers from myostatin-deficient mice are associated with a suppression of protein degradation.</a> J Appl Physiol. 111 (1), 185-191 (2011).</li><li>Gumucio, J. P., Davis, M. E., Bradley, J. R., Stafford, P. L., Schiffman, C. J., Lynch, E. B., Claflin, D. R., Bedi, A., Mendias, C. 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Assessments of the contractile properties of permeabilized single skeletal muscle fibers are used to investigate muscle function in a wide variety of contexts. Examples include studies that have evaluated the effects of aging12, exercise10,13,14, spaceflight15, injury2,3,16, drug treatments17,18, disease19 and genetic manipulation20,21 on fiber structure and function. Due to the ability to directly assess the contractile performance of myofibrils in their native configuration, this technique provides an attractive platform from which to form an understanding of myofibrillar function absent of potentially confounding effects that are present when neuromuscular signal transmission and excitation-induced calcium release are included in the system under study. Furthermore, functional testing of single fibers can be used to complement contractile protein identification results such as those obtained through immunohistochemistry or gel electrophoresis + western blot22.
One of the primary functions of skeletal muscle is to generate force. Consequently sFo, a measure of the intrinsic force generating capability of a contractile system, is of great interest to muscle physiologists. Reliable estimates of sFo require accurate measures of both fiber CSA and Fo. Since fibers are, in general, neither circular in cross-section, nor uniform in CSA along their length, great care should be taken when estimating CSA. To this end, measurements are made at several locations along the length of the fiber and, at each location, from two perspectives separated by 90&#176;. Reliable measures of Fo require attention to several details including accounting for passive force, adjusting sarcomere length to maximize overlap of thick and thin filaments, employing an activating solution with a calcium concentration that results in maximal activation, maintaining the desired experimental temperature, and maintaining optimal storage conditions (temperature and duration) of the fibers prior the day of the experiment.
While the steps outlined here describe the procedure for evaluating maximum isometric force, it is frequently desirable to evaluate other important functional qualities of skeletal muscle fibers. This can be achieved by extending the experimental protocol to include additional mechanical manipulations of the fiber. For example, measurement of the speed at which the fiber shortens against a series of different loads allows determination of the force-velocity relationship, from which force-power and velocity-power relationships can be computed10,23,24. Additionally, the speed of unloaded shortening can be determined by employing the &#8220;slack test&#8221;25, which consist of applying a series of slack-inducing shortening steps and measuring the time required by the fiber to remove the slack. Another kinetic parameter that is frequently reported is ktr, the rate constant for force redevelopment following a mechanical perturbation that temporarily detaches all crossbridges26. Finally, the relationship between calcium concentration and active force generation (the &#8220;force-pCa relationship&#8221;) is often of interest18 and can be determined by exposing the fiber to a series of solutions with calcium concentrations ranging from below the threshold for activating the contractile system to those sufficient to elicit maximum activation and therefore maximum force (Fo).
Though much of the mentioned equipment is needed for assessing single fiber contractility, other equipment is not absolutely necessary. The length-controller, for example, is essential for any experimental protocol that requires rapid or precise lengthening or shortening of the fiber, but is not absolutely necessary for evaluating maximum isometric force (though a zero-force level in the force record must still be identified by some means). The prisms that allow observation of the fiber from the side, while useful for assessing cross-sectional area, are not absolutely necessary when positioning the fiber within the experimental chamber. Furthermore, alternative means for exposing the fiber to the various experimental solutions could be employed, including devising a manually-operated system of chambers or a single chamber that allows for rapid filling and emptying of solutions. Finally, while sub-physiological experimental temperatures such as 15 &#176;C are commonly used to improve the reproducibility of mechanical measurements1,2,3,5,8,12,17,27, it is possible to generate valid data at other temperatures23,28 as long as the effects of temperature on solution properties (calcium concentration, pH, etc.) are taken into consideration.
The compositions of the testing solutions are among the most critical aspects of the permeabilized fiber techniques described here. Considerations regarding solution composition are complex and beyond the scope of this article. The solutions described in Step 5 of the protocol section are designed with an emphasis on rapid activation of the permeabilized fiber upon its transfer from pre-activating to activating solutions while maintaining a constant ionic strength, cationic composition, and osmolarity6,29. Other approaches to solution composition have been employed with notable success by other research groups and typically make use of published binding constants and computational tools27,30,31. The concentrations of calcium ions in the various activating solutions is particularly important in studies involving submaximal activation such as force-pCa evaluations. For experiments in which fibers are fully activated, such as those described here, the calcium concentration in the activating solution typically exceeds by a comfortable margin that required to achieve maximum force, making its precise knowledge less critical. Addition of creatine phosphate is important for buffering the intramyofibrillar ATP and ADP fluctuations that would otherwise be associated with contractile activity. Creatine kinase is required to catalyze the phosphate transfer from creatine phosphate to ADP. Under experimental conditions that result in high ATP turnover rates, including working at high temperatures or measuring high-speed shortening in fast fibers32, creatine kinase must be added to the solution to supplement the endogenous creatine kinase that remains bound to the fiber. For less demanding experimental conditions, the ATP regeneration system is less critical27.
Limitations of the permeabilized single fiber technique include the following. The data generated by these tests define the contractile properties of the specific myofibrillar unit that was attached to the experimental apparatus. Consequently, this captures only a small fraction of the entire multinucleated fiber from which the segment was obtained which, in turn, represents a small fraction of the total number of fibers within the muscle. Investigators should thus consider carefully the sampling required to support any conclusions drawn from the experiments. Additionally, evaluating the impact of an exercise training intervention on fiber function presumes that the fibers evaluated were indeed recruited during the training. Though the protocol attempts to mimic the natural intracellular milieu of the fiber, the sarcolemma permeabilization process is non-specific and necessarily allows soluble intracellular constituents to freely diffuse into the bathing solutions. A further consequence of the membrane permeability is a change in the osmotic balance evidenced by a swelling in fiber volume33. The fiber swelling increases the distance between actin and myosin filaments resulting in reduced calcium sensitivity of the myofilament system34,35, but can be reversed by the introduction of large, osmotically active compounds34. A final limitation to consider is the consequence of the technique used to attach fibers to the experimental apparatus. This invariably requires distorting the spatial relationship within the filament system at and near the attachment points, with attending functional deficits. Specifically, the regions of the fiber at and adjacent to the attachment points are functionally compromised and thereby contribute artifactual series elasticity to the measurement system.
In summary, we have described a means by which to assess the force-generating capacity of chemically permeabilized skeletal muscle fibers in vitro. Though the focus of this article has been on the assessment of maximum isometric force generating capacity of human skeletal muscle fibers, the experimental approach can be modified and extended to determine a variety of kinetic parameters and relationships across a range of species, mammalian or otherwise.

The primary function of skeletal muscle is to generate force. Muscle force is elicited in vivo through a complex sequence of events that includes motor nerve action potentials, neuromuscular transmission, muscle fiber action potentials, release of intracellular calcium, and activation of the system of regulatory and contractile proteins. Because force generation is the ultimate result of this sequence, a deficit in force could be caused by failure of one or more of the individual steps. A key attribute of the permeabilized fiber preparation is that it eliminates most of the steps required for force generation in vivo, with only the regulatory and contractile functions associated with the myofibrillar apparatus remaining. The investigator assumes control over the delivery of activating calcium and energy (ATP), and is rewarded with a simplified system that allows assessment of the isolated regulatory and contractile structures in their native configuration. Measurements of force using permeabilized skeletal muscle fibers are thus valuable when assessing alterations in muscle function observed in vivo. For example, we have used this technique to characterize the force generating capacity of fibers from myostatin deficient mice1 and to assess the cause of persistent muscle weakness exhibited following chronic rotator cuff tears2,3.
Modern permeabilized fiber methodology can be traced to early influential studies4,5 and is currently in use by a number of research groups. Though the techniques have been described in the literature, they have not yet been presented in video format. The goal of this article is to illustrate an updated, valid and reliable technique for measuring the maximum force generating capacity of single fibers from chemically permeabilized skeletal muscle samples. To accomplish this, an individual fiber segment (referred to herein as a &#8220;fiber&#8221;) is extracted from a pre-permeabilized bundle of fibers and placed in an experimental chamber containing a relaxing solution, the defining feature of which is a calcium concentration that is &#60;10 nM. The fiber is then attached at one end to a force-transducer and at the other end to a length-controller. With the fiber held at an optimal sarcomere length, it is transferred to an activating solution that has a calcium concentration sufficient to elicit maximum activation and thereby maximum isometric contraction force. Force data are acquired, stored and analyzed using a personal computer.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Polystyrene culture test tube with cap</td>
			<td>Fisher Scientific&#160;</td>
			<td>14-956-3D</td>
			<td></td>
		</tr>
		<tr>
			<td>0.5 mL screw cap micocentrifuge</td>
			<td>Fisher Scientific&#160;</td>
			<td>02-681-334</td>
			<td></td>
		</tr>
		<tr>
			<td>0.5 mL microcentrifuge caps with o-ring</td>
			<td>Fisher Scientific&#160;</td>
			<td>02-681-358</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge cryobox</td>
			<td>Fisher Scientific&#160;</td>
			<td>5055-5005</td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Mettler-Toledo</td>
			<td>FE20</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Fisher Scientific&#160;</td>
			<td>08-757-11YZ</td>
			<td></td>
		</tr>
		<tr>
			<td>Nonsterile-suture 10-0 monofilament</td>
			<td>Ashaway Line Twine</td>
			<td>S30002</td>
			<td></td>
		</tr>
		<tr>
			<td>Insect pins</td>
			<td>Fine Science Tools</td>
			<td>26002-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps - Dumont #5</td>
			<td>Fine Science Tools</td>
			<td>11251-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Microdissecting scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td>Leica Microsystems</td>
			<td>MZ8</td>
			<td></td>
		</tr>
		<tr>
			<td>Micrometer drives</td>
			<td>Parker Hannifin</td>
			<td>3936M</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermometer</td>
			<td>Physitemp</td>
			<td>BAT-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Water bath circulator&#160;</td>
			<td>Neslab Instruments</td>
			<td>RTE-111</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature controller</td>
			<td>Aplha Omega Instruments</td>
			<td>Series 800</td>
			<td></td>
		</tr>
		<tr>
			<td>LabVIEW software</td>
			<td>National Instruments</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Computer</td>
			<td>Varied</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Chamber system</td>
			<td>Aurora Scientific</td>
			<td>802D</td>
			<td></td>
		</tr>
		<tr>
			<td>Length-controller</td>
			<td>Aurora Scientific</td>
			<td>312C</td>
			<td></td>
		</tr>
		<tr>
			<td>Force-transducer</td>
			<td>Aurora Scientific</td>
			<td>403A</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagents</td>
		</tr>
		<tr>
			<td>K-proprionate</td>
			<td>TCI America</td>
			<td>P0510</td>
			<td></td>
		</tr>
		<tr>
			<td>Imadizole</td>
			<td>Sigma-Aldrich</td>
			<td>I0125</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2&#8226;6H20</td>
			<td>Sigma-Aldrich</td>
			<td>M2670</td>
			<td></td>
		</tr>
		<tr>
			<td>Brij 58</td>
			<td>Sigma-Aldrich</td>
			<td>P5884</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA (acid)</td>
			<td>Sigma-Aldrich</td>
			<td>E0396</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2H2ATP&#8226;0.56H2O</td>
			<td>Sigma-Aldrich</td>
			<td>A7699</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma-Aldrich</td>
			<td>G6279</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (acid)</td>
			<td>Sigma-Aldrich</td>
			<td>H7523</td>
			<td></td>
		</tr>
		<tr>
			<td>MgO</td>
			<td>Sigma-Aldrich</td>
			<td>529699</td>
			<td></td>
		</tr>
		<tr>
			<td>HDTA (acid)</td>
			<td>TCI America</td>
			<td>D2019</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCO3</td>
			<td>Sigma-Aldrich</td>
			<td>C4830</td>
			<td></td>
		</tr>
		<tr>
			<td>NaN3</td>
			<td>Sigma-Aldrich</td>
			<td>S8032</td>
			<td></td>
		</tr>
		<tr>
			<td>KOH (1N)</td>
			<td>Sigma-Aldrich</td>
			<td>35113</td>
			<td></td>
		</tr>
		<tr>
			<td>HCL (1N)</td>
			<td>Sigma-Aldrich</td>
			<td>318949</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2CrP&#8226;4H2O</td>
			<td>Sigma-Aldrich</td>
			<td>P7936</td>
			<td></td>
		</tr>
		<tr>
			<td>pH 10 standard</td>
			<td>Fisher Scientific</td>
			<td>SB115</td>
			<td></td>
		</tr>
		<tr>
			<td>pH 7 standard</td>
			<td>Fisher Scientific</td>
			<td>SB107</td>
			<td></td>
		</tr>
	</tbody>
</table>

measurement of maximum isometric force generated by permeabilized skeletal muscle fibers

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. Single muscle fiber studies are useful in both basic science and clinical studies. For basic studies, single muscle fiber contractility measurements allow investigation of fundamental mechanisms of force production, and analysis of muscle function in the context of genetic manipulations. Clinically, single muscle fiber studies provide useful insight into the impact of injury and disease on muscle function, and may be used to guide the understanding of muscular pathologies. In this video article we outline the steps required to prepare and isolate an individual skeletal muscle fiber segment, attach it to force-measuring apparatus, activate it to produce maximum isometric force, and estimate its cross-sectional area for the purpose of normalizing the force produced.

Healthy, chemically permeabilized single fibers should appear uniform in shape and have consistent striation spacing when viewed under high magnification. Fibers that are inflexible when manipulated with the forceps or have obvious structural damage should be discarded.
High magnification digital images taken during step 15 are analyzed for 5 paired diameter measurements along the midsection of the fiber. Fiber CSA is estimated assuming an elliptical cross section and averaging 5 individual CSA measurements as depicted in Figure 7A. Figure 7B also serves to illustrate how fiber dimensions in one view can be significantly different compared with paired dimensions in the other view (i.e., cross-sections are not, in general, round).
Representative force traces from human slow and fast fibers are shown in Figures 8A and 8B, respectively. Voltage output of the force-transducer is acquired during a test and converted to force (mN) using data acquisition and analysis software (LabVIEW). Figure 9 illustrates the approach used to assess maximum active force (Fo), which is calculated by subtracting the force required to maintain the fiber at optimal sarcomere length while in a relaxed state (passive force, FP), from the greatest isometric force developed during maximal fiber activation (total force, FT). Since the output of the force-transducer that corresponds to zero force is, in general, different for each of the different bathing chambers, we briefly slacken the fiber in both the pre-activating and activating solutions to capture the zero-force level in the experimental record. Normalization of maximum active force by fiber CSA is used to generate the more informative value of specific force (sFo). Because it takes into consideration the CSA of the fiber, sFo provides a measure of the intrinsic force generating capacity of the fiber&#8217;s contractile apparatus, thereby allowing functional comparisons between fibers of disparate sizes. It should, however, be noted that CSA measurements are not able to distinguish the proportion of the fiber occupied by contractile filaments versus the proportion occupied by other subcellular structures.
Typical characteristics of healthy, adult fibers from Claflin et al. 201110 for human, Mendias et al. 20111 for mouse and Gumucio et al. 20122 for rat are detailed in Table 3. All data presented in Table 3 were generated using the techniques described in this article.
<table border="0" cellpadding="0" cellspacing="0" style="text-align:center">
	<tbody>
		<tr>
			<td rowspan="3" style="border:0px #fff solid"></td>
			<td colspan="4">Human 
			(vastus lateralis)</td>
			<td>Mouse 
			(EDL)</td>
			<td nowrap="nowrap">Rat 
			(infraspinatus)</td>
		</tr>
		<tr>
			<td colspan="2">Male</td>
			<td colspan="2">Female</td>
			<td>Male</td>
			<td>Male</td>
		</tr>
		<tr>
			<td>Type 1</td>
			<td>Type 2a</td>
			<td>Type 1</td>
			<td>Type 2a</td>
			<td>(not typed)</td>
			<td>(not typed)</td>
		</tr>
		<tr>
			<td>CSA (&#956;m2)</td>
			<td>4880 - 6900</td>
			<td>5270 - 8380</td>
			<td>3870 - 5470</td>
			<td>4010 - 5610</td>
			<td>1850 - 3080</td>
			<td>5290 - 8010</td>
		</tr>
		<tr>
			<td>Fo (mN)</td>
			<td>0.79 - 1.17</td>
			<td>1.02 - 1.54</td>
			<td>0.64 - 0.97</td>
			<td>0.71 - 1.07</td>
			<td>0.14 - 0.25</td>
			<td>0.55 - 0.97</td>
		</tr>
		<tr>
			<td>sFo (kPa)</td>
			<td>142 - 182</td>
			<td>165 - 210</td>
			<td>156 - 193</td>
			<td>172 - 214</td>
			<td>67 - 94</td>
			<td>102 - 131</td>
		</tr>
		<tr>
			<td>n</td>
			<td>129</td>
			<td>160</td>
			<td>149</td>
			<td>207</td>
			<td>37</td>
			<td>94</td>
		</tr>
	</tbody>
</table>

Table 3. Typical characteristics of healthy, adult fibers from human vastus lateralis10, mouse extensor digitorum longus1 and rat infraspinatus2 muscles. Optimal sarcomere lengths were set at 2.7 &#181;m for human fibers7,8 and 2.5 &#181;m for both mouse (36,37) and rat fibers (38). Experimental Lf ranges (25th and 75th quartiles) were 1.39-1.73 mm, 1.17-1.53 mm and 1.32-1.59 mm for human, mouse and rat respectively. Ranges shown indicate the 25th and 75th quartiles and n is the number of fibers tested.
The most common problems experienced during testing include a suture loop slip, which results in a force response with a &#8220;catch&#8221; such as that illustrated in Figure 10A, and a partial or full thickness tear of the fiber, which results in a force response that returns abruptly toward or to (break) zero while the fiber is still immersed in activating solution (Figure 10B). If a slip, tear or break occurs during an experiment, the fiber should be discarded and the data excluded, though maintaining a record of fiber failures can also be informative11. Another negative outcome that may be encountered is the premature activation of the fiber while in the pre-activating solution (Figure 10C). Partial activation in the pre-activating solution suggests significant cross-well contamination (i.e., an unintentional increase in calcium concentration in the pre-activating well). In this instance, all baths should be aspirated and rinsed well with deionized water. Drying the dividing surfaces between the chambers is also recommended as moisture or condensation in these areas may lead to wicking of solution between baths. The decision to include or exclude data will ultimately depend on the experimental focus and should thus be considered when designing the study.
<img alt="Figure 1" src="/files/ftp_upload/52695/52695fig1.jpg" /> 
Figure 1: Suture loop (10-0 monofilament nylon suture).
<img alt="Figure 2" src="/files/ftp_upload/52695/52695fig2.jpg" /> 
Figure 2: Bundle dissection. Forceps are in left hand, microdissection scissors are in right hand. Red line indicates the favorable orientation of the wrist and scissors with the longitudinal axes of the fibers.
<img alt="Figure 3" src="/files/ftp_upload/52695/52695fig3.jpg" /> 
Figure 3: (A) Testing apparatus with labeled components. (a) Experimental chambers with transparent bottoms. (b) Length-controller. (c) Force-transducer. (d) Light source. (e) Length-controller x-y-z micrometer drive with digital display. (f) Stage micrometer drive with digital display. (g) Force-transducer x-y-z micrometer drive. (h) Platform for calibrated laser-diffraction target screen. (i) Vibration isolation table. (B) Close-up view of the experimental chambers. (j) Stainless steel attachment surface extending from the length-controller. (k) Stainless steel attachment surface extending from the force-transducer. (l) Side-view prism. (m) Housing for thermoelectric cooling modules. (n) Thermocouple for reporting chamber temperature.
<img alt="Figure 4" src="/files/ftp_upload/52695/52695fig4.jpg" /> 
Figure 4: Modified 100 &#181;l pipet tip used to transfer fiber from dissection dish to experimental chamber.
<img alt="Figure 5" src="/files/ftp_upload/52695/52695fig5.jpg" /> 
Figure 5: Mounting single fiber onto experimental apparatus. (A) Prepared suture loops threaded onto stainless steel attachment surfaces. (B) Fiber transferred to experimental chamber. (C) Fiber anchored to stainless steel attachment surfaces by first pair of suture loops with excess suture removed. (D) Second pair of suture loops threaded over the top of first suture loops and tied in place.
<img alt="Figure 6" src="/files/ftp_upload/52695/52695fig6.jpg" /> 
Figure 6: Sarcomere length is assessed by the projection of a laser interference pattern onto a calibrated target screen. (a) Laser source. (b) Mirror. (c) Target screen. (d) Laser interference pattern.
<img alt="Figure 7A" src="/files/ftp_upload/52695/52695fig7A.jpg" />
<img alt="Figure 7B" src="/files/ftp_upload/52695/52695fig7B.jpg" /> 
Figure 7: (A) Determination of fiber cross-sectional area at optimal sarcomere length (human = 2.7 &#181;m). Assuming an elliptical cross-section, CSA is calculated for each of five locations along the fiber midsection and the mean of the five individual measurements is reported as fiber CSA. 2a represents top view diameter and is one axis of the ellipse, 2b represents side view diameter and is the other axis of the ellipse. (B) Representative fiber images illustrating each of the five corresponding diameter measurements taken in both the top and side view.
<img alt="Figure 8" src="/files/ftp_upload/52695/52695fig8.jpg" /> 
Figure 8: Representative force traces from healthy human vastus lateralis muscle fibers. (A) Type 1 fiber (CSA: 5710 &#181;m2, Fo: 0.89 mN and sFo: 156 kPa). (B) Type 2a fiber (CSA: 9510 &#181;m2, Fo: 1.66 mN and sFo: 174 kPa). Fiber myosin heavy chain type was determined through the use of electrophoretic separation and silver-staining techniques22.
<img alt="Figure 9" src="/files/ftp_upload/52695/52695fig9.jpg" /> 
Figure 9: Calculation of maximum active force (Fo). (a) Expanded view of fiber force response during slack-inducing movement of length-controller initiated in pre-activating solution. FP is the force required to maintain a sarcomere length of 2.7 &#181;m with the fiber at rest. (b) Expanded view of length-controller slack-inducing movement. Note that Fo = FT&#8211; FP.
<img alt="Figure 10" src="/files/ftp_upload/52695/52695fig10.jpg" /> 
Figure 10: (A) Suture loop slip, evidenced by a &#8220;catch&#8221; in force trace during the rise of force. Check to be sure loops are secure before activating the fiber. (B) Fiber break during activation. May be due to poor fiber integrity or aggressive fiber treatment during suture loop placement. (C) Premature partial fiber activation due to contamination of pre-activation chamber with Ca2+.

Watch this Scientific Journal Video about Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers at JoVE.com

Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. In this article we outline a valid and reliable technique to prepare and test permeabilized skeletal muscle fibers in vitro.

Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle ...

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Bioengineering

25.1K Views.  University of Michigan Medical School. Analysis of the contractile properties of chemically skinned, or permeabilized, skeletal muscle fibers offers a powerful means by which to assess muscle function at the level of the single muscle cell. In this article we outline a valid and reliable technique to prepare and test permeabilized skeletal muscle fibers in vitro.

Video: Measurement of Maximum Isometric Force Generated by Permeabilized Skeletal Muscle Fibers

Measurement of Aggregate Cohesion by Tissue Surface Tensiometry

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium. We have developed tissue surface tensiometry (TST) specifically to measure the surface free energy of interaction between cells. The biophysical concepts underlying TST have been previously described in detail1,2. The method is based on the observation that mutually cohesive cells, if maintained in shaking culture, will spontaneously assemble into clusters. Over time, these clusters will round up to form spheres. This rounding-up behavior mimics the behavior characteristic of liquid systems. Intercellular binding energy is measured by compressing spherical aggregates between parallel plates in a custom-designed tissue surface tensiometer. The same mathematical equation used to measure the surface tension of a liquid droplet is used to measure surface tension of 3D tissue-like spherical aggregates. The cellular equivalent of liquid surface tension is intercellular binding energy, or more generally, tissue cohesivity. Previous studies from our laboratory have shown that tissue surface tension (1) predicts how two groups of embryonic cells will interact with one another1-5, (2) can strongly influence the ability of tissues to interact with biomaterials6, (3) can be altered not only through direct manipulation of cadherin-based intercellular cohesion7, but also by manipulation of key ECM molecules such as FN8-11 and 4) correlates with invasive potential of lung cancer12, fibrosarcoma13, brain tumor14 and prostate tumor cell lines15. In this article we will describe the apparatus, detail the steps required to generate spheroids, to load the spheroids into the tensiometer chamber, to initiate aggregate compression, and to analyze and validate the tissue surface tension measurements generated.

We describe a method of measuring binding energy, expressible as tissue surface tension, between cells within 3D tissue-like aggregates. Differences in tissue surface tension have been demonstrated to correlate with invasiveness of lung, muscle, and brain tumors, and are fundamental determinants of establishing spatial relationships between different cell types.

Rigorous measurement of intercellular binding energy can only be made using methods grounded in thermodynamic principles in systems at equilibrium.  We ...

Bacterial Immobilization for Imaging by Atomic Force Microscopy

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface. It has the ability to image cells and biomolecules, in a liquid environment, without the need to chemically treat the sample. In order to accomplish this goal, the sample must sufficiently adhere to the mounting surface to prevent removal by forces exerted by the scanning AFM cantilever tip. In many instances, successful imaging depends on immobilization of the sample to the mounting surface. Optimally, immobilization should be minimally invasive to the sample such that metabolic processes and functional attributes are not compromised. By coating freshly cleaved mica surfaces with porcine (pig) gelatin, negatively charged bacteria can be immobilized on the surface and imaged in liquid by AFM. Immobilization of bacterial cells on gelatin-coated mica is most likely due to electrostatic interaction between the negatively charged bacteria and the positively charged gelatin. Several factors can interfere with bacterial immobilization, including chemical constituents of the liquid in which the bacteria are suspended, the incubation time of the bacteria on the gelatin coated mica, surface characteristics of the bacterial strain and the medium in which the bacteria are imaged. Overall, the use of gelatin-coated mica is found to be generally applicable for imaging microbial cells.

Live Gram-negative and Gram-positive bacteria can be immobilized on gelatin-coated mica and imaged in liquid using Atomic Force Microscopy (AFM).

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface.  It has the ability to image cells and biomolecules, in a liquid ...

Engineering Skeletal Muscle Tissues from Murine Myoblast Progenitor Cells and Application of Electrical Stimulation

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to study pressure ulcers, for regenerative medicine and as a meat alternative 1. The first reported 3D muscle constructs have been made many years ago and pioneers in the field are Vandenburgh and colleagues 2,3. Advances made in muscle tissue engineering are not only the result from the vast gain in knowledge of biochemical factors, stem cells and progenitor cells, but are in particular based on insights gained by researchers that physical factors play essential roles in the control of cell behavior and tissue development. State-of-the-art engineered muscle constructs currently consist of cell-populated hydrogel constructs. In our lab these generally consist of murine myoblast progenitor cells, isolated from murine hind limb muscles or a murine myoblast cell line C2C12, mixed with a mixture of collagen/Matrigel and plated between two anchoring points, mimicking the muscle ligaments. Other cells may be considered as well, e.g. alternative cell lines such as L6 rat myoblasts 4, neonatal muscle derived progenitor cells 5, cells derived from adult muscle tissues from other species such as human 6 or even induced pluripotent stem cells (iPS cells) 7. Cell contractility causes alignment of the cells along the long axis of the construct 8,9 and differentiation of the muscle progenitor cells after approximately one week of culture. Moreover, the application of electrical stimulation can enhance the process of differentiation to some extent 8. Because of its limited size (8 x 2 x 0.5 mm) the complete tissue can be analyzed using confocal microscopy to monitor e.g. viability, differentiation and cell alignment. Depending on the specific application the requirements for the engineered muscle tissue will vary; e.g. use for regenerative medicine requires the up scaling of tissue size and vascularization, while to serve as a meat alternative translation to other species is necessary.

Engineered muscle tissue has great potential in regenerative medicine, as disease model and also as an alternative source for meat. Here we describe the engineering of a muscle construct, in this case from mouse myoblast progenitor cells, and the stimulation by electrical pulses.

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to ...

Biosensor for Detection of Antibiotic Resistant Staphylococcus Bacteria

A structurally transformed lytic bacteriophage having a broad host range of Staphylococcus aureus strains and a penicillin-binding protein (PBP 2a) antibody conjugated latex beads have been utilized to create a biosensor designed for discrimination of methicillin resistant (MRSA) and sensitive (MSSA) S. aureus species 1,2. The lytic phages have been converted into phage spheroids by contact with water-chloroform interface. Phage spheroid monolayers have been moved onto a biosensor surface by Langmuir-Blodgett (LB) technique 3. The created biosensors have been examined by a quartz crystal microbalance with dissipation tracking (QCM-D) to evaluate bacteria-phage interactions. Bacteria-spheroid interactions led to reduced resonance frequency and a rise in dissipation energy for both MRSA and MSSA strains. After the bacterial binding, these sensors have been further exposed to the penicillin-binding protein antibody latex beads. Sensors analyzed with MRSA responded to PBP 2a antibody beads; although sensors inspected with MSSA gave no response. This experimental distinction determines an unambiguous discrimination between methicillin resistant and sensitive S. aureus strains. Equally bound and unbound bacteriophages suppress bacterial growth on surfaces and in water suspensions. Once lytic phages are changed into spheroids, they retain their strong lytic activity and show high bacterial capture capability. The phage and phage spheroids can be utilized for testing and sterilization of antibiotic resistant microorganisms. Other applications may include use in bacteriophage therapy and antimicrobial surfaces.

Lytic phage biosensors and antibody beads are able to discriminate between methicillin resistant (MRSA) and sensitive staphylococcus bacteria. The phages were immobilized by a Langmuir-Blodgett method onto a surface of a quartz crystal microbalance sensor and worked as broad range staphylococcus probes. Antibody beads recognize MRSA.

A structurally transformed lytic bacteriophage having a broad host range of Staphylococcus aureus strains and a penicillin-binding protein (PBP 2a) ...

Preparation of Light-responsive Membranes by a Combined Surface Grafting and Postmodification Process

In order to modify the surface tension of commercial available track-edged polymer membranes, a procedure of surface-initiated polymerization is presented. The polymerization from the membrane surface is induced by plasma treatment of the membrane, followed by reacting the membrane surface with a methanolic solution of 2-hydroxyethyl methacrylate (HEMA). Special attention is given to the process parameters for the plasma treatment prior to the polymerization on the surface. For example, the influence of the plasma-treatment on different types of membranes (e.g.&#160;polyester, polycarbonate, polyvinylidene fluoride) is studied. Furthermore, the time-dependent stability of the surface-grafted membranes is shown by contact angle measurements. When grafting poly(2-hydroxyethyl methacrylate) (PHEMA) in this way, the surface can be further modified by esterification of the alcohol moiety of the polymer with a carboxylic acid function of the desired substance. These reactions can therefore be used for the functionalization of the membrane surface. For example, the surface tension of the membrane can be changed or a desired functionality as the presented light-responsiveness can be inserted. This is demonstrated by reacting PHEMA with a carboxylic acid functionalized spirobenzopyran unit which leads to a light-responsive membrane. The choice of solvent plays a major role in the postmodification step and is discussed in more detail in this paper. The permeability measurements of such functionalized membranes are performed using a Franz cell with an external light source. By changing the wavelength of the light from the visible to the UV-range, a change of permeability of aqueous caffeine solutions is observed.

A plasma-induced polymerization procedure is described for the surface-initiated polymerization on polymer membranes. Further postmodification of the grafted polymer with photochromic substances is presented with a protocol of conducting permeability measurements of light-responsive membranes.

In order to modify the surface tension of commercial available track-edged polymer membranes, a procedure of surface-initiated polymerization is ...

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. Kelvin probe force microscopy (KPFM) is a module of atomic force microscopy (AFM) that measures the contact potential difference between surfaces at the nano-scale. The combination of KPFM with AFM allows for the simultaneous generation of surface potential and topographical maps of biological samples such as bacterial cells. Here, we employ KPFM to examine the effects of surface potential on microbial adhesion to medically relevant surfaces such as stainless steel and gold. Surface potential maps revealed differences in surface potential for microbial membranes on different material substrates. A step-height graph was generated to show the difference in surface potential at a boundary area between the substrate surface and microorganisms. Changes in cellular membrane surface potential have been linked with changes in cellular metabolism and motility. Therefore, KPFM represents a powerful tool that can be utilized to examine the changes of microbial membrane surface potential upon adhesion to various substrate surfaces. In this study, we demonstrate the procedure to characterize the surface potential of individual methicillin-resistant Staphylococcus aureus USA100 cells on stainless steel and gold using KPFM.

Here, we present a protocol explaining the use of Kelvin probe force microscopy as a tool for generating high resolution nano-scale surface potential maps. This tool was applied to assess the role of surface potential on the binding capacity of microorganisms to substrate surfaces.

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. ...

Detection of Exosomal Biomarker by Electric Field-induced Release and Measurement (EFIRM)

Exosomes are microvesicular structures that play a mediating role in intercellular communication. It is of interest to study the internal cargo of exosomes to determine if they carry disease discriminatory biomarkers. For performing exosomal analysis, it is necessary to develop a method for extracting and analyzing exosomes from target biofluids without damaging the internal content.
Electric field-induced release and measurement (EFIRM) is a method for specifically extracting exosomes from biofluids, unloading their cargo, and testing their internal RNA/protein content. Using an anti-human CD63 specific antibody magnetic microparticle, exosomes are first precipitated from biofluids. Following extraction, low-voltage electric cyclic square waves (CSW) are applied to disrupt the vesicular membrane and cause cargo unloading. The content of the exosome is hybridized to DNA primers or antibodies immobilized on an electrode surface for quantification of molecular content.
The EFIRM method is advantageous for extraction of exosomes and unloading cargo for analysis without lysis buffer. This method is capable of performing specific detection of both RNA and protein biomarker targets in the exosome. EFIRM extracts exosomes specifically based on their surface markers as opposed to size-based techniques.
Transmission electron microscopy (TEM) and assay demonstrate the functionality of the method for exosome capture and analysis. The EFIRM method was applied to exosomal analysis of 9 mice injected with human lung cancer H640 cells (a cell line transfected to express the exosome marker human CD63-GFP) in order to test their exosome profile against 11 mice receiving saline controls. Elevated levels of exosomal biomarkers (reference gene GAPDH and protein surface marker human CD63-GFP) were found for the H640 injected mice in both serum and saliva samples. Furthermore, saliva and serum samples were demonstrated to have linearity (R = 0.79). These results are suggestive for the viability of salivary exosome biomarkers for detection of distal diseases.

Detection of Exosomal Biomarker by Electric Field-induced Release and Measurement EFIRM

Exosomes are microvesicular structures found within biofluids that potentially carry important disease discriminatory biomarkers. Here, a novel method is used to specifically extract exosomes and rapidly test the exosomal cargo for both RNA/protein targets following the disruption of exosomes using non-uniform electric cyclic square waves.

Exosomes are microvesicular structures that play a mediating role in intercellular communication. It is of interest to study the internal cargo of ...

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from traumatic injury, disease and various congenital, genetic and acquired conditions are quite common. Tissue engineering and regenerative medicine technologies have enormous potential to provide a therapeutic solution. However, utilization of biologically relevant animal models in combination with longitudinal assessments of pertinent functional measures are critical to the development of improved regenerative therapeutics for treatment of VML-like injuries. In that regard, a commercial muscle lever system can be used to measure length, tension, force and velocity parameters in skeletal muscle. We used this system, in conjunction with a high power, bi-phase stimulator, to measure in vivo force production in response to activation of the anterior crural compartment of the rat hindlimb. We have previously used this equipment to assess the functional impact of VML injury on the tibialis anterior (TA) muscle, as well as the extent of functional recovery following treatment of the injured TA muscle with our tissue engineered muscle repair (TEMR) technology. For such studies, the left foot of an anaesthetized rat is securely anchored to a footplate linked to a servomotor, and the common peroneal nerve is stimulated by two percutaneous needle electrodes to elicit muscle contraction and dorsiflexion of the foot. The peroneal nerve stimulation-induced muscle contraction is measured over a range of stimulation frequencies (1-200 Hz), to ensure an eventual plateau in force production that allows for an accurate determination of peak tetanic force. In addition to evaluation of the extent of VML injury as well as the degree of functional recovery following treatment, this methodology can be easily applied to study diverse aspects of muscle physiology and pathophysiology. Such an approach should assist with the more rational development of improved therapeutics for muscle repair and regeneration.

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

We describe an in vivo protocol to measure dorsiflexion of the foot following stimulation of the peroneal nerve and contraction of the anterior crural compartment of the rat hindlimb. Such measurements are an indispensable translational tool for evaluating skeletal muscle pathology and tissue engineering approaches to muscle repair and regeneration.

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.