extfieldsofstudy: Medicine Biology

text:
Expression of tropomyosin in relation to myofibrillogenesis in axolotl hearts

The anatomy, function and embryonic development of the heart have been of interest to clinicians and researchers alike for centuries. A beating heart is one of the key criteria in defining life or death in humans. An understanding of the multitude of genetic and functional elements that interplay to form such a complex organ is slowly evolving with new genetic, molecular and experimental techniques. Despite the need for ever more complex molecular techniques some of our biggest leaps in knowledge come from nature itself through observations of mutations that create natural defects in function. Such a natural mutation is found in the Mexican axolotl, Ambystoma mexicanum. It is a facultative neotenous salamander well studied for its ability to regenerate severed limbs and tail. Interestingly it also well suited to studying segmental heart development and differential sarcomere protein expression due to a naturally occurring mendelian recessive mutation in cardiac mutant gene “c”. The resultant mutants are identified by their failure to beat and can be studied for extended periods before they finally die due to lack of circulation. Studies have shown a differential expression of tropomyosin between the conus and the ventricle indicating two different cardiac segments. Tropomyosin protein, but not its transcript have been found to be deficient in mutant ventricles and sarcomere formation can be rescued by the addition of TM protein or cDNA. Although once thought to be due to endoderm induction our findings indicate a translational regulatory mechanism that may ultimately control the level of tropomyosin protein in axolotl hearts.

Introduction
The Mexican axolotl, a facultative neotenous salamander, provides a valuable model to study heart development due to a cardiac lethal mutation (gene c) that affects only heart muscle [1,2]. It has also been used extensively for organ regeneration research, particularly of its limbs and tails [3] but have included initial studies into the regeneration of the heart [4]. The axolotl cardiac gene c mutation is a Mendelian, autosomal recessive lethal mutation with significant effects on tropomyosin protein levels in the cardiac tissue. Morphological studies of the abnormal cardiomyogenesis in mutants have shown they lack organized myofibrils, have large collections of amorphous material, but still retain normal electrophysiological properties [5][6][7]. The embryos can survive for up to a fortnight post-hatching which is ideal for studying this process before the lack of circulation, secondary to abnormal sarcomere formation is lethal. The mutant axolotl heart provides a unique opportunity for studying the intricate process of cardiac development and for examining the specific functional role of each tropomyosin (TM) isoform in this process. Ultimately the protein level of TM is profoundly diminished in the ventricle of c/c mutant hearts, resulting in an absence of organized myofibrils and subsequently the inability to beat [5][6][7][8].
It is important to note that the conus is not deficient in tropomyosin protein, retains organized myofibrils and is capable of beating independently, unlike the atria and ventricle in mutant hearts [8]. Notably, the mutant hearts can be rescued in situ by supplying exogenous TM protein or TM cDNA in an expression construct under the control of an appropriate promoter(s) [9,10]. Mutant hearts can also be rescued in situ by a specific non-coding RNA that is unrelated to TM [11][12][13].
However, the exact mechanism by which this RNA modulates the expression of tropomyosin is yet to be elucidated. To better understand the mechanism(s) effecting tropomyosin expression in mutant hearts, we undertook an extensive molecular characterization of the various isoforms of tropomyosin in the Mexican axolotl.

Isoform diversity of tropomyosin in vertebrates
The thin filaments of striated muscle in vertebrate consist of actin, tropomyosin, the troponin (Tn) complex (Tn-I, Tn-C and Tn-T), tropomodulin, and a few other proteins [14]. Actin filaments interaction with Ca +2 governs muscle contraction and relaxation. Tropomyosin is a coiled coil actin-binding protein found along the length of seven actin monomers. A set of four known genes (TPM1, TPM2, TPM3 and TPM4) that encode tropomyosin in vertebrates [15][16][17][18][19] give rise to various tropomyosin protein isoforms that play important roles in striated, smooth and non-muscle cells. There remains even further diversity in other models such as zebrafish where six tropomyosin genes have been identified [15]. The creation of different tropomyosin isoforms occurs through various mechanisms, including the use of different promoters, alternative mRNA splicing, different 3' end mRNA processing and tissue specific translation control [20].
We have cloned and sequenced the cDNA of three sarcomeric TM isoforms from cardiac tissues. These isoforms are designated as TPM1α, TPM1κ, and TPM4α [21][22][23] (Table 1). TPM1α, one of nine alternatively spliced isoforms of the TPM1 gene, is known to be the major sarcomeric isoform in mammalian hearts [15][16][17][18][19]. We first identified and characterized another alternatively spliced sarcomeric isoform of the TPM1 gene in axolotl hearts [22], designated TPM1κ. TPM1α and TPM1κ have an identical exon composition except for exon 2 where TPM1κ contains exon 2a instead of exon 2b ( Figure 1 and Table 1). Exon 2a is characteristic of the smooth muscle type isoform (TPM1β) of the TPM1 gene ( Figure 1 and Table 1). TPM1κ transcripts and its corresponding protein are expressed in both axolotl hearts and skeletal muscle there appears to be a differential translation of the transcript. Using qRT-PCR the expression level of TPM1κ transcripts is higher than TPM1α (ratio α:κ is 0.32) in adult axolotl hearts although TPM1κ protein is less than 10% of the total sarcomeric TM, as determined by CH1 antibody [24]. The opposite is true in adult skeletal muscle where the level of TPM1κ transcript is significantly lower compared to TPM1α (ratio of α:κ > 13) but the level of TPM1κ protein constitutes~30% of the total sarcomeric TM [24] Similarly, the levels of expression of TPM1α and TPM1κ transcripts in human hearts are comparable but the actual TPM1κ protein level is only~5% of the total sarcomeric TM [25]. Comparatively, TPM1α protein con-stitutes~90-95% of the total TM in human hearts [24,25] while TPM1κ is not expressed in human skeletal muscle at all [26]. TPM1κ transcripts are also expressed in embryonic chicken heart but not in adult heart and skeletal muscle [27]. It remains unknown if the protein is expressed since human TPM1κ antibody may not cross-react with chicken TPM1κ protein [25]. This discrepancy between transcript and protein levels in the two isoforms in heart tissue suggests that TPM1κ transcripts may undergo translational repression.
Accounting for the other tropomyosin genes other than TPM1, TPM2α (sarcomeric isoform of the TPM2 gene) is also expressed in mammalian hearts in addition to the previously described TPM1α and TPM1κ The sarcomeric isoform of the TPM3 gene, TPM3α, is only expressed in slow-twitch skeletal muscle. No sarcomeric isoform of the TPM4 gene is expressed in mammalian striated muscles because the TPM4 gene is truncated in mammals [15][16][17]. On the contrary, TPM4α is a major TM isoform in amphibian cardiac tissues [23,31] and is the only isoform for sarcomeric TM in adult avian hearts [27,30,32].

Sarcomeric tm protein in cardiac mutant axolotl hearts
Among the various myofibril proteins, tropomyosin has been shown by a variety of experiments to be drastically reduced in cardiac mutant hearts [2,6,9,12]. Increasing the intracellular levels of TM in cardiac mutant heart cells via introduction of FITC-labeled exogenous TM protein by itself or an expression construct allowing in-vivo TM production subsequently promoted myofibrillogenesis ( Figure 2) [9]. Control mutant heart stained with CH1 monoclonal antibody specific for sarcomeric TM, demonstrated minimal staining when examined by confocal microscopy ( Figure 2b). However, examination of mutant heart transfected with an expression construct of murine TPM1α cDNA under the control of mouse α-MYHC promoter demonstrated the formation of organized myofibrils ( Figure 2c). The results prove mutant hearts are capable of forming cardiac myofibrils when provided with sufficient levels of tropomyosin protein. This unequivocally demonstrated the functional defect in the gene "c" mutation is the deficiency in tropomyosin protein although the underlying cause of this functional deficit is less clear. Interestingly, other myofibril structural proteins such as actin, myosin and myosin binding protein C (MyBp-C) were found to be at or near normal levels in the mutant hearts [5,28] while one protein, tropomodulin which is intricately related functionally with sarcomere maturation is increased [33]. The most comprehensive study on the analysis of sarcomeric tropomyosin protein expression in normal and mutant axolotl hearts were reported by Zhang et al. [12]. To determine whether multiple isoforms of tropomyosin exist in embryonic axolotl hearts and to verify if they are differentially regulated in mutant hearts, 2D western blot with the monoclonal antibody (CH1), was performed. Five different protein spots (tropomyosin isoforms) from both normal and mutant embryonic hearts at stages 36 to 42 were detected ( Figure 3). All isoforms of tropomyosin detected by the sarcomer specific CH1 Figure 1 Alternative splicing patterns of TPM1 and TPM4 genes. (a) Exon composition of the TPM1 gene and alternative splicing that generates two striated muscle specific isoforms -TPM1α and TPM1κ [26]. (b) Exon composition of TPM4 gene (adapted from avian species as proposed by Fleenor et al. [30] and alternative splicing generate two isoforms in axolotl, TPM4α and a low-molecular weight TPM4 transcript. antibody (30) were located between pI 4 to 5 with molecular weight of~38 kD. The results showed protein levels of the 4 CH1 recognized TM isoforms were decreased significantly in mutant hearts compared to normal hearts (Figure 3a and 3b) [12].
Although mutant axolotl hearts are deficient in sarcomere specific TM proteins, mRNA levels of each of three striated muscle isoforms (TPM1α, TPM1κ, and TPM4α) are comparable in normal and mutant hearts [12,23]. Hence, the tropomyosin deficiency in mutant heart is not due to an insufficiency in transcription or post-transcriptional splicing [23]. We cloned and sequenced cDNAs of three isoforms from mutant hearts; no mutation(s) was detected in any of these cDNAs that may cause truncated non-functional TM isoform(s). Additionally, we have cloned and sequenced the promoter region of the TPM4 gene from the DNA isolated from normal and mutant axolotl hearts and again, no differences were observed [34]. Hence, the possibility of insufficient transcription of the cardiac specific TPM4α isoform is highly unlikely. The most plausible explanation based on available evidence of TM deficiency in mutant hearts is a translational insufficiency of the tropomyosin transcripts in mutant hearts [12].

Molecular analysis and manipulation of tropomyosin isoforms in normal and mutant axolotl hearts
As stated earlier, there are at least three striated muscle isoforms of tropomyosin present in the axolotl. Two isoforms of tropomyosin cDNA have been identified which apparently are derived from the single alpha-tropomyosin gene (TPM1) through alternative splicing [21,22]. Spinner et al. [23] cloned another tropomyosin cDNA, which is the product of a TM4 type tropomyosin gene from axolotl heart. An expression construct with each of these isoforms upon transfection into mutant hearts canaugment tropomyosin proteinlevels and promotes myofibrillogenesis. The important question is whether or not any one of these  c. Stage-36 mutant heart lipofected with an expression construct containing a murine TPM1α cDNA under the control of α-Myosin Heavy Chain promoter, which subsequently induced TM and promoted myofibrillogenesis. Mutant heart stained with α-actinin primary antibody. Well-organized sarcomeric myofibrils can be seen throughout the heart (arrows). Staining of the Z-lines confirmed the sarcomeric organization seen in TPM1α transfected hearts that were stained with tropomyosin primary antibody (results not shown here) [ref]. Interestingly, statge-36 mutant heart tansfected with a murine TPM2α cDNA under the same promoter did express some TM protein but sarcomeric myofibrils did not form throughout the heart in contrast to TPM1α transfected hearts [7]. isoforms alone and/or in various combination(s) is necessary for myofibrllogenesis in axolotl hearts in vivo. In order to address this issue we developed procedures for disruption of myofibrils in normal axolotl hearts mimicking the mutant hearts by lipofecting antibodies against sarcomeric TM into normal hearts in situ. Myofibrils in lipofected normal hearts indeed became greatly disorganized [10,35]. As CH1 antibodies react with all three sarcomeric tropomyosins, it is not possible to evaluate the requirement of a particular isoform that is involved in cardiac myofibrillogenesis. Later we developed antibody against TPM1κ in rabbits using a 15-mer peptide sequence (LDELHKSEESLLTAD) derived from axolotl exon 2a [36]. Recall, TPM1κ is unique as a sarcomeric TM in that it contains exon 2a instead of exon 2b which is found in TPM1α. The affinity purified anti-TPM1κ antibody upon transfection could disarray the organized myofibrils in axolotl hearts [37]. The results strongly suggest that TPM1κ plays a critical role in myofibril formation in axolotl hearts.
Additionally, isoform specific sense and anti-sense oligonucleotide was transfected into normal axolotl hearts. TPM1κ expression was blocked in whole embryonic axolotl heart by transfection of exon 2a-specific anti-sense oligonucleotide (Figure 4b). In contrast, myofibrils were unaffected in normal control heart when transfected with FITC label sense oligonucleotides (Figure 4c). RNA was isolated from treated and untreated hearts and subsequently RT-PCR was carried out with isoform specific primer-pairs. The results confirmed the lower Figure 4 Transfection of isoform-specific sense and anti-sense oligonucleotides in the ventricle of normal hearts. a. Confocal microscopy of normal axolotl hearts transfected with TPM1a anti-sense oligonucleoted and subsequently stained with CH1 monoclonal antibody. TPM1a anti-sense oligonucleotide did not result in a drastic disruption of organized myofibrils in the ventricle, which is comparable with the normal untreated control hearts (figure not shown). Sarcomeric TM can be seen in most of the cells. Contractility of the anti-sense treated hearts were not affected. b. TPM1κ anti-sense transfection disrupted the myofibril organization in normal axolotl heart compared to TPM1κ sense transfection. Very little organized structure is seen when examined by the tropomyosin staining. The secondary antibody is contained within amorphous areas in the cells. c. It shows that the TPM1κ sense oligonucleoides did not affect the structure. Since no effect on myofibril structure with TPM1κ sense was observed, we tagged the oligonucleotide with FITC (green) to verify its presence and found it to be within the myocytes. Double staining of the nucleus (green) and the myofibrils at the periphery of the cells (red) can be seen. The green staining is ovoid in shape and primarily located at the center of the cells [10]. Confocal z-series images of stage~38 embryonic axolotl hearts transfected with either TPM1κ anti-sense or sense oligonucleotides. Immunodetection of sarcomeric tropomyosin using CH1 monoclonal antibody is shown in red. The results depicted by this figure suggests that TPM1κ plays a critical role in maintaining the myofibrillar structure in embryonic axolotl hearts.
transcript expression of TPM1κ in anti-sense treated hearts. The conclusion was substantiated by the in vitro analysis of the specificity of the TPM1κ anti-sense oligonucleotides used in this study. Confocal analysis of the sense and anti-sense oligonucleotide transfected normal axolotl hearts was carried out after staining with antitropomyosin antibody (CH1). Immunohistochemical analysis unequivocally confirmed that the inhibition of the expression of TPM1κ disrupted myofibril structure of the myofibrils in anti-sense transfected normal axolotl hearts. In contrast, TPM1α anti-sense oligonucleotide did not cause a disruption of the myofibrillar organization in axolotl hearts (Figure 4a).
In a separate study, we found that the antisense TPM4α oligonucleotide disrupted myofibril formation and inhibited beating in normal axolotl hearts, while the sense strands did not. A fluorescein-tagged sense oligonucleotide clearly showed that the oligonucleotide was introduced within the cells of intact hearts. The results implicate the essential role of TPM4α in cardiac myofibrillogenesis ( Figure 5) [8,10,36].

Differential expression of tropomyosin in conus and the ventricle
Despite the dramatic and lethal effects of the homozygous cardiac gene "c" mutation in the axolotl ventricle, the conus of the heart beats and has organized myofibrils ( Figure 6). In order to understand whether various Figure 5 Effect of transfection of TM isoform-specific sense-and anti-sense oligonucleotides on myofibril organization in hearts from normal axolotl hearts. a. Stereo anaglyph of a 24 section confocal laser scanning microscope z-series of normal axolotl heart stained with CH1 anti-tropomyosin antibody. This type of image is shown because it demonstrates branching myofibrils in whole hearts. b. Stereo anaglyph of normal hearts transfected with TPM4α exon 2-specific anti-sense, 5′-T*A*C*T*AGCTCGTCCTCAAGC*T*G*C*-3′, where N* represents the phophothioate blocked oligonucleotide. Myofibirl organization was disrupted in a majority of the cardiomyocytes. Most of the tropomyosin appears to be in amorphous areas when expressed. Some of the cells do not appear to have a detectable level of tropomyosin. Gross morphology was normal and the cells appeared to be intact although myofibril structure was largely disrupted. The contractility was diminished significantly [36]. c. Normal heart transfected with TPM4α exon 2-specific sense chimeric olighonucleotide, 5′-fG*C*A*GCTTGAAGGCGAGC TA*G*T*A*-3, where *N represents phosphothioate blocked nucleotide, and fG represents G tagged with Fluorescein at the 5′end. This image is a compressed z-series of 2 sections that was not stereo offset but used to demonstrate double staining. Isolated pieces of myofibrils that were contained within these sections of the cardiac cells were stained with tropomyosin antibody (re, arrowheads). This image is primarily useful for demonstrating the presence green fluorescence (GFP) within the nuclear area of the cariomyocytes by five days (asterik). Hoescht staining of the nuclei in the same heart coincided with the FITC staining that were localized in a majority of nuclei (figure not shown). The figure was adapted from Spinner et al. (2004) [36]. TM isoforms are differentially expressed in different segments of the heart and whether the known TM isoforms contribute to myofibril formation in a segment specific manner, we employed anti-sense oligonucleotides to separately knockdown post-transcriptional expression of TPM1α and TPM4α in axolotl heart segments. We evaluated the organization of myofibrils in the conus and ventricle of normal and cardiac mutant hearts using immunohistochemical techniques. We concluded that the TPM1α isoform, a product of the TPM1 gene, was essential for myofibrillogenesis in the conus, whereas TPM4α, the striated muscle isoform of the TPM4 gene, was essential for myofibrillogenesis in the ventricle. Our results support the segmental theory of vertebrate heart development and suggest the conus is a different transcriptional tissue unit. Since the conus is an outflow tract structure and in humans is a conical pouch of the right ventricle from which the pulmonary artery arises it will be interesting to examine tropomyosin isoform diversity in these two segments in other systems including humans [8]. Development of the conus appears to be unaffected in the mutant heart and is comparable to the normal heart. The different functions of the heart segments would suggest that different isoforms could be needed in accordance with that function. The physiologic characteristics necessary for the ventricle versus the conus and subsequent outflow tract are different [38]. Isoform diversity in specific heart segments includes tropomyosin but also could include other sarcomeric proteins such as myosin heavy chain [39] Further studies could examine the localization and function of specific isoforms in adult heart segments. There has also been research on isoform diversity and the relationship to diseases such as dilated cardiomyopathy [25]. These studies suggest that with changing physiologic parameters, the isoforms can also be changed. Ultimately, the study of segment specific tropomyosin isoforms may help in the understanding of time and function specific sarcomeric proteins and their relationship to regeneration of heart function in damaged hearts.

Promotion of myofibrillogenesis in mutant hearts in situ by a non-coding rna
A noncoding RNA, Myofibril-Inducing RNA (MIR) is capable of promoting myofibrillogenesis and heart beating in the mutant (c/c) axolotl hearts in situ [11]. Zhang et al. [12] demonstrated that the MIR gene is essential for tropomyosin (TM) expression in axolotl hearts during development at the level of translation or posttranslation. qRT-PCR using isoform-specific primer-pairs showed that mRNA expression of three sarcomeric tropomyosin isoforms (TPM1α, TPM1κ, and TPM4α) in untreated mutant hearts and in normal hearts knocked down with double-stranded MIR (dsMIR) are similar to untreated normal. However, at the protein level, sarcomeric tropomyosin isoforms detected with CH1 monoclonal antibodies, are significantly reduced in mutant and dsMIR treated normal hearts. However, this study neither showed the mechanism by which MIR may induce sarcomeric TM synthesis in axolotl hearts nor addressed the role of specific tropomyosin isoforms in cardiac myofibrillogenesis.
Recently, Kochegarovr et al. [40] randomly cloned RNAs from fetal human heart. RNA from one of the clones (clone #291) was found to promote myofibril formation in mutant axolotl in situ. This RNA induced expression of cardiac markers in mutant hearts: tropomyosin, troponin and αsyntrophin. The nucleotide sequences of the cloned RNA matches in partial with the human microRNA-499a and b, although it differs in length. qRT-PCR data suggest this RNA may induce the TPM4α (ATmC-3) isoform in mutant heart, producing more sarcomeric tropomyosin protein and subsequently promote myofibril formation. The mechanism by which this non-coding RNA induces tropomyosin in mutant hearts may well be different from that of MIR, which acts at the post transcriptional level [12].

Review and conclusions
Although our complete understanding of the mechanism of tropomyosin expression in mutant axolotl hearts as well as the nature and function of gene "c" is far from over, we would like to end this review with a positive note. Our finding of sarcomeric TM isoform TPM1κ in axolotl led to the discovery of this isoform in human hearts [26]. Unlike in axolotl, it is not expressed in human skeletal muscle. Most importantly, an upregulation of TPM1κ protein has been reported in hearts from human dilated cardiomyopathy patients [25]. However, it is not yet known whether the upregulation of TPM1κ is the cause or a consequence of cardiomyopathy in this (See figure on previous page.) Figure 6 Tropmodulin but not tropomyosin found in cardiac "c" mutant axolotl ventricle. CH1 anti-sarcomeric TM antibody labels the conus but not the ventricle in stage 38/39 cardiac mutant axolotl hearts. Stage 38/39 normal (a-c) and mutant (d-f) axolotl hearts were double stained for immunofluorescent microscopy with CH1 anti-sarcomeric TM antibody (FITC channel) and polyclonal anti-tropomodulin antibody, R1749 (Rhodamine channel). Only the FITC channel is shown for whole heart images (a, d) demonstrating the lack of staining in the ventricle of mutant hearts while all higher magnification images utilize a dual rhodamine-FITC filter. Letter labeled boxes in A and D correspond with the area shown in their respective higher magnification image. TM labeled with CH1 antibody is found in a cross-striated pattern in the ventricle of normal hearts (c) and in the conus of both normal (b) and mutant (e) hearts. Tropomyosin staining is markedly absent in mutant ventricle (f) resulting in a failure of normal sarcomere formation with tropmodulin found in linear arrays near the surface of the membranes (Arrows, F). Scale Bar = 100 μm. patient. Our anti-sense experiments suggest strongly the functional significance of TPM1κ in axolotl hearts. In addition, the lower expression level of TPM1κ protein in axolotl heart and skeletal muscle [24] and also in human hearts [25] points towards translational repression of TPM1κ. Further, upon injection intraperitoneally into juvenile axolotl, Shz-1, a cardiogenic small molecule, augmented the expression levels of transcripts of TPM1α, TPM1κ, and TPM4α in hearts. But the increased transcript level did not resulted into increased sarcomeric TM protein expression [41]. Finally, although the transcript levels of all TPM isoforms in normal and mutant axolotl heart ventricles are comparable, the proteins of all three isoforms are diminished significantly. This observation also point towards the translational repression of TM in cardiac tissues [42]. The evidence for translational control of sarcomeric TM in mammalian hearts was originally came from the works from the laboratory of Dr. David Wieczorek, University of Cincinnati, Cincinnati, OH [29,43]. Rethinesamy et al. [29] and Blanchard et al. [44] independently ablated one of the two alleles of the TPM1 gene in mice that resulted in half of the TPM1α transcripts in ablated mice hearts compared to wild-type. However, TPM1α protein level was unchanged in ablated mice hearts suggesting a higher translational efficiency of TPM1α transcripts in ablated mice hearts. Again, Rajan et al. [25] reported that although the level of transcripts of TPM1α and TPM1κ human hearts is parallel, TPM1κ protein is only~5% of the total sarcomeric TM whereas TPM1α protein constitutes about 90-95% of the total sarcomeric TM. The results strongly suggest the translational repression of TPM1κ transcripts in human hearts. The immediate future goal of our laboratory is to explore further the translational repression of tropomyosin expression in vertebrate hearts as well as to find out the functional role of TPM1κ.


extfieldsofstudy: Biology Medicine

text:
Divergent RNA transcription

New approaches using biotinylated-psoralen as a probe for investigating DNA structure have revealed new insights into the relationship between DNA supercoiling, transcription and chromatin compaction. We explore a hypothesis that divergent RNA transcription generates negative supercoiling at promoters facilitating initiation complex formation and subsequent promoter clearance.

N ew approaches using biotinylatedpsoralen as a probe for investigating DNA structure have revealed new insights into the relationship between DNA supercoiling, transcription and chromatin compaction. We explore a hypothesis that divergent RNA transcription generates negative supercoiling at promoters facilitating initiation complex formation and subsequent promoter clearance.
Packaging DNA into chromatin creates a repressive environment. This is thought to reduce transcriptional noise and provide additional levels of gene regulation. Many processes have been described that alter chromatin architecture and facilitate or repress transcription. These include transcription factor binding, recruitment of RNA polymerases, ATP-dependent chromatin remodeling and changes in histone modifications. The level of DNA supercoiling is also known to be important for transcription but remains poorly understood, as few techniques are available for directly investigating it. This is about to change: new approaches have recently been used for analyzing DNA topology in cells [1][2][3] and show transcription dependent changes in DNA supercoiling that impact on higher levels of chromatin structure. Furthermore, new data indicates that DNA supercoiling and divergent RNA transcription are linked providing new ideas for understanding the role of DNA topology in regulating gene transcription.
The structure of DNA within mammalian cells is not known, although it is often thought to be in a B-form configuration. However, this will depend on the base

Divergent RNA transcription
A role in promoter unwinding? Catherine Naughton, Samuel Corless and Nick Gilbert* MRC Human Genetics Unit; Institute of Genetics and Molecular Medicine; The University of Edinburgh; Edinburgh, UK composition, extent of supercoiling 4 and DNA binding proteins, such as HMGs. As DNA moves through the RNA polymerase positive (over-wound) supercoils are generated ahead of the polymerase and negative (under-wound) supercoils behind, described as the twin supercoiled domain model. 5 Therefore, transcription generates significant torsion in the DNA, so the observation that topoisomerase I, an enzyme that relaxes supercoils, is associated with the transcription machinery 6 and localizes to actively transcribed chromatin 7 led to the suggestion that it might relieve topological strain. However, early experiments show that genes are more efficiently transcribed if they are encoded on negatively supercoiled plasmids. 8,9 In addition, superhelical tension is also a prerequisite for TFIID binding 10 promoting the formation of a preinitiation complex, 11 transcription factor binding 4 and promoter clearance. 12 Consequently, cell based experiments using psoralen, as a probe for DNA supercoiling, show that promoter regions are under-wound 1,13 and supercoiling may prime specific genes for transcription. 14 Together this suggests that DNA supercoiling must be precisely controlled and is an additional level of transcriptional regulation. However, how DNA supercoiling is introduced at TSS prior to transcription remains unanswered.
A major clue arose from an in vivo study demonstrating that local domains of supercoiling can affect transcription. 15 In this work, a promoter is activated due to transcription generated negative supercoils from a nearby divergent promoter and inhibition of endogenous topoisomerase I further increases transcription. While this ( Fig. 2C). Our data suggests that the initiating form of RNA polymerase, functioning both upstream through divergent transcription and downstream through abortive sense transcription, generates short transcripts and negatively supercoils TSSs creating a permissive environment for subsequent transcription. Furthermore, as DNA supercoiling impacts on higher levels of chromatin organization, 1 there is a possible spreading of supercoiling from one locus to another, which could facilitate transcription of surrounding genes and might provide a rationale for gene clustering in the human genome.
This model harmonizes recent advances in our understanding of transcriptional regulation. Traditionally, RNAP II recruitment is thought to be the rate limiting step and, thus, the key regulatory step in eukaryotic transcription; however, genome wide profiling of RNAP II indicates that it is bound and initiated at both active and inactive genes. 25 Indeed, for a large proportion of metazoan genes (20-30%), RNAP II density is enriched downstream of many TSSs and this has been described as RNAP II promoter proximal pausing. 26,27 Pausing is now thought to be a widespread regulatory mechanism with the Negative Elongation Factor (NELF) and DRB-Sensitivity Inducing Factor (DSIF) protein complexes binding to and arresting RNAP II 60 nts downstream of the TSS. Subsequent recruitment of Positive Transcription Elongation Factor b (P-TEFb) to this paused RNAP II complex and phosposphorylation of DSIF, NELF and Ser2 on the RNAP II C-terminal results in dissociation of NELF and productive transcriptional elongation. Both sense and antisense RNAP II complexes are involved in RNAP II pausing and both depend on PTEF-b recruitment. 19 The purpose of pausing is not known but it is frequently found at developmental control genes and stimulus-responsive pathways and is though to allow their rapid and synchronous induction in response to extracellular signals. 27 Consequently, loss of pausing through knockdown of the pause-inducing factor NELF leads to broadly attenuated immune gene activation. 28 One function of paused RNAP II is to establish a permissive chromatin environment 29 and paused polymerase has its under-wound level. Treatment of cells with bleomycin (introducing DNA nicks) relaxes DNA supercoiling demonstrating that promoter regions are under topological tension, which is also regulated by topoisomerase I and II. 1,2 As transcription clearly introduced negative DNA supercoils under-winding promoter regions, we investigated further the role of divergent initiation in this process. In our experimental approach, the elongating form of RNA polymerase II (RNAP II) is very sensitive to α-amanitin and is rapidly degraded in a proteasome dependent manner (Fig. 1B). In contrast, the initiating form of the polymerase is more resistant to degradation suggesting that changes in supercoiling upon α-amanitin washout are manifest by the initiating form of the polymerase. This was additionally confirmed as blocking transcription elongation specifically with flavopiridol rapidly reduced long RNA production, but there was a lag before a drop in the synthesis of short RNAs, consistent with initiating RNA polymerase producing short RNAs. 1 Thus, we reasoned that elongating polymerase synthesizes long RNAs while the initiating form produces shorter RNA species. 24 To test this, we trace labeled cells with 3H-Uridine and showed that the production of long RNAs (> 200 nt) was substantially reduced after 5 h α-amanitin treatment and continued to decrease after α-amanitin washout (Fig. 1C). In contrast, short RNAs (< 200 nt) were produced in abundance by 1 h recovery and continued to increase at 2 h (Fig. 1C), concomitant with remodeling of DNA supercoiling (Fig. 1A). We hypothesized that these short RNAs are the products of divergent transcription. [16][17][18] Using data from Core et al., 16 we investigated the expression of several short antisense or sense RNA transcripts around the promoters of the expressed IGBP1 (Xq13.1) and LDHA (11p15.1) genes ( Fig. 2A).
Long RNAs produced within the gene body were inhibited by α-amanitin and did not recover within 2 h (Fig. 2B). However, in agreement to what is seen globally (Fig. 1C), short antisense and sense RNAs are produced upstream and downstream of genes a short period of time after α-amanitin washout and recovery study exploited in vitro generated plasmid templates, divergent transcription, whereby transcription is initiated in both sense and anti-sense directions from a promoter, has been shown to be a feature of many active mammalian genes. [16][17][18] The high resolution mapping of short RNA transcripts reveals extensive sense and anti-sense transcript peaks at approximately 50 bp and 250 bp respectively from the TSS. Antisense RNAs are capped 19 but are present at a relatively low abundance as they are substrates for the exosome. 18 Histone modifications reflective of transcription initiation (H3K4me3) are found both in the sense and antisense direction while histone marks indicative of transcriptional elongation (H3K79me2) are found in the sense direction. 17,20 Consequently divergent transcription is more appropriately described as divergent initiation as only productive elongation occurs in the direction of the gene.
Divergent initiation is very prevalent (approximately 80% of transcribed mammalian genes) but does it have a regulatory function or can it just represent "sloppy" transcription initiation events resulting from an open promoter architecture? Several functions have been proposed 21 but, in particular, Seila et al. 22 suggest divergent initiation may under-wind promoter regions facilitating productive transcription initiation and elongation. To better investigate DNA supercoiling we have developed an approach for directly analyzing it in cells. 1 This technique builds on previous studies that have exploited the cell permeable drug tri-methyl psoralen (TMP), which preferentially intercalates and can be cross-linked into negatively supercoiled (under-wound) DNA. 23 The addition of a biotin group, attached via a linker, to the psoralen (bTMP) enables the selective enrichment of under-wound DNA fragments that can be mapped by hybridization to high-resolution genomic microarrays. In RPE1 cells, bTMP binding analysis demonstrates that TSSs are under topological strain and negatively supercoiled in a region that extends ~20 kb into the body of the gene and 10 kb upstream (Fig. 1A). Transcription inhibition by α-amanitin or DRB 2 causes significant remodeling of TSS DNA to a more positively supercoiled state and subsequent α-amanitin washout promotes a remodeling of the TSS back to yet been assigned to the short sense and antisense RNAs further substantiates our idea that they are by products of a critical process necessary to create a transcriptionally friendly chromatin environment. Future work combining targeted RNAi of RNAP II pausing and elongation factors with DNA supercoiling analysis will elucidate the mechanism by which chromatin structure influences transcription.

Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
with developmentally regulated genes, while constitutively expressed genes are maintained with optimal levels of DNA supercoiling. However, anti-sense RNA transcription is significantly less prevalent in Drosophila, 24,31 but there is pronounced promoter pausing. This may suggest that the purpose of pausing may be different in mammalian and Drosophila genomes, or that abortive sense transcription is sufficient to negatively supercoil the promoter to facilitate transcription factor binding and subsequent processive transcription. The fact that no function has as been shown to block nucleosome assembly at promoters, thus maintaining an open chromatin architecture. 20,30 In support of this, we propose that promoter proximal pausing permits time for divergent transcription to produce short RNAs and, concomitantly, negatively supercoil promoter regions to facilitate transcription. After transcriptional pause-release, polymerases in collaboration with topoisomerase and helicases can then maintain the supercoiling state of the locus in a regulated manner. This is consistent with pausing being more frequently associated Cells were suspended in 2 x SDS lysis buffer, incubated at 100°C for 5 min and sonicated briefly. Protein samples were resolved on 8% bis-tris gels and transferred to PVDF membrane by wet-transfer. Membranes were probed with antibodies using standard techniques and detected by enhanced cheiluminscence. RNA polymerase II antibodies: initiating RNA polymerase H14, 1:500 (Covance, MMS-134R) and elongating RNA polymerase H5, 1:500 (Covance, MMS-129R). GAPDH, 1:1000 (Cell Signaling, #2118). (C) Graph showing 30 min pulsed incorporation of 185 nM [5-3 H] Uridine into short and long RNA species after transcription inhibition. Cold dA, dG, dC, dT and C (37 nM final) were added to cells, to suppress label incorporation into DNA. After 30 min incubation cells were rinsed with PBS and long and short RNAs was extracted by selective binding to a silica matrix (miRNeasy Kit, Qiagen). Residual DNA was removed by on-column DNaseI treatment. RNA was quantified using a Nanodrop and 3 H incorporation was measured by scintillation counting. were detected by first poly adenylating and then reverse transcribed using tagged oligo-dT and random primers (miScript kit, Qiagen). They were then quantified using qPCR with a specific forward primer and universal reverse primer (miScript primer assay, Qiagen). Primer sequences are: IGBP1 −277 TTGTCTCTCT ACCGCCTTCC IGBP1 −17 GAAGATCCGG TCGCTTGAG LDHA +193 CGATTCCGGA TCTCATTG LDHA +279 AGGGATGGGC GGGTAGAG.


extfieldsofstudy: Physics Medicine

text:
Muscle residual force enhancement: a brief review

Muscle residual force enhancement has been observed in different muscle preparations for more than half a century. Nonetheless, its mechanism remains unclear; to date, there are three generally accepted hypotheses: 1) sarcomere length non-uniformity, 2) engagement of passive elements, and 3) an increased number of cross-bridges. The first hypothesis uses sarcomere non-homogeneity and instability to explain how “weak” sarcomeres would convey the higher tension generated by an enhanced overlap from “stronger” sarcomeres, allowing the whole system to produce higher forces than predicted by the force-length relationship; non-uniformity provides theoretical support for a large amount of the experimental data. The second hypothesis suggests that passive elements within the sarcomeres (i.e., titin) could gain strain upon calcium activation followed by stretch. Finally, the third hypothesis suggests that muscle stretch after activation would alter cross-bridge kinetics to increase the number of attached cross-bridges. Presently, we cannot completely rule out any of the three hypotheses. Different experimental results suggest that the mechanisms on which these three hypotheses are based could all coexist.

& INTRODUCTION
When a muscle is stretched while activated and held at a certain final length long enough for the force transients to cease, the steady force achieved is always higher than the steady force that develops when the muscle is activated while already held isometrically at the same final length (1-5) ( Figure 1). This residual force enhancement was first observed by Abbott & Aubert (1) and predated Huxley's cross-bridge model (6). Nevertheless, this enhancement could not be accounted for by either this model (6,7) or the force-length relationship (8); to date, the mechanism of force enhancement remains elusive.
Additionally, it is decreased but not fully abolished after deactivation (10,18).
Several possible explanations for residual force enhancement have been proposed, although none are able to fully elucidate its mechanism (3). This paper provides a brief review of the three generally accepted hypotheses for residual force enhancement: 1) sarcomere length nonuniformity, 2) engagement of a passive element, and 3) increase in the number of cross-bridges. Our prime goal is to recapitulate and summarize these three hypotheses for residual force enhancement in a more digestible manner, allowing a wider audience to understand a phenomenon that can also be relevant to other fields, such as exercise physiology, biomechanics, healthy-gait mechanics, physical therapy, etc. If more in-depth information is needed, we suggest the following reviews to cover specific aspects of residual force enhancement: Herzog et al. (3), Rassier and Herzog (20), Edman (21), and Rassier (22).

Hypothesis 1 -Sarcomere length non-uniformity
Sarcomere length non-uniformity was the first mechanism used to explain force enhancement after stretch (23), being still currently accepted and advocated (21). The first study to present a possible explanation and indirect evidence that sarcomere length non-uniformity was involved in residual force enhancement was performed by Julian and Morgan (24). Later, Morgan (25) provided a more complete explanation that could account for most of the residual force enhancement characteristics found experimentally. One of the bases for non-uniformity within sarcomeres is the phenomenon of instability on the descending limb of the force-length relationship (26). An unstable system allows differences in individual sarcomere lengths: some sarcomeres elongate more, becoming weaker (due to decreasing filament overlap), and others in turn become even stronger (due to increasing overlap). The ''weaker'' sarcomeres continue to elongate until a critical point at which they ''yield;'' in other words, the ''weaker'' sarcomeres' cross-bridges can no longer maintain adequate tension, which leads to their detachment (yield). These sarcomeres continue to elongate until the tension borne by passive (non-contractile) elements equals the tension of the ''strong'' sarcomeres, and in this case, the ''weak'' sarcomeres are ''popped.'' The remaining tensile force is now determined by the stronger sarcomeres, which ''transmit'' their tension through the ''popped'' sarcomeres, thereby enabling the entire system to exhibit greater force than would be predicted by the average sarcomere force based on the force-length relationship curve (25).

Support for the theory
Several studies (1,24,27) have shown that force enhancement occurs only on the descending limb of the force-length curve, which is believed to be unstable (26). In support of this idea, Morgan (25) showed that in the absence of instability, sarcomeres would theoretically stretch uniformly, and no enhancement should be expected.
Sarcomere length non-uniformities may also provide an explanation for different force-enhancement behaviors under differing conditions, such as velocity independence of force enhancement and stretch-magnitude dependence (1,2). Muscle-stretch velocity neither prevents nor affects instability (25,26); therefore, sarcomere length non-uniformity must be independent from stretch velocity. In contrast, force enhancement should be directly related to stretch magnitude because the more a muscle is stretched, the greater the number and degree of sarcomere elongation and greater the production of residual tension. A mathematical explanation has been provided for this behavior (25). What Julian and Morgan (24) noted in their study was that sarcomeres from the centers of the cells stretched more than sarcomeres near the ends. Talbot and Morgan (28) confirmed this observation. Using electron microscopy, they were able to observe that certain half-sarcomeres were more elongated than the average half-sarcomere.
One of the strongest pieces of evidence presented by sarcomere-non-uniformity advocates is the decrease in stiffness. Assuming that the number of formed crossbridges is the main factor responsible for increased muscle stiffness, both longer and ''popped'' sarcomeres should each present a lower stiffness than the shorter sarcomeres. Therefore, stiffness should decrease after a muscle is stretched. This expected decrease in stiffness has been reported in some studies (29)(30)(31).

Limitations and criticisms
The idea of instability has long been one of the bases for explaining sarcomere length non-uniformity. This notion was explored by Morgan (25) from Hill's (26) idea of dynamic instability derived from the negative slope on the descending limb of the force-length curve. Conceptually, this approach was inadequate because one cannot assume dynamic instability from a construct based on observations made under isometric conditions (3). However, even when the sarcomeres were clamped and kept stable to prevent non-uniformities, force enhancement was still observed (1). More recently, results obtained using myofibril preparations showed that although sarcomeres did not stretch uniformly on either the descending (32) or ascending (4) limb of the force-length curve, they remained stable after stretch, indicating that force enhancement can occur even in the absence of instability.
A number of studies have shown force enhancement on the supposedly stable ascending limb of the force-length curve. These studies were performed not only in fibers (13,33) but also in muscles in situ (9,10,18) and even in myofibrils (4). All of these studies reported small but consistent force enhancements on the ascending limb.
According to the sarcomere length non-uniformity theory (25), force enhancement after stretch should not be higher than the tension produced at the initial length, let alone exceed the force at the optimal sarcomere length (plateau). Again, several experiments performed with different muscle preparations have refuted this assumption and shown that force enhancement could exceed the maximal isometric force developed at the plateau of the force-length relationship (4,10,13,15,(17)(18)(19).
In contrast to studies that have found decreased stiffness in the force-enhanced state (29)(30)(31), other studies have found an increase in stiffness when comparing the isometric reference force with the force in the enhanced state in different muscle preparations (34,35). Clearly, controversy remains regarding the explanation for stiffness behavior after stretch.
Finally, the development of techniques that have enabled the investigation of isolated myofibrils (36) and even single sarcomeres (37) has brought further skepticism regarding the sarcomere length non-uniformity theory. Studies have found force enhancement not only while sarcomere lengths were kept stable within activated myofibrils that were stretched (4,15) but also in isolated single sarcomeres (17). Both studies rule out sarcomere length non-uniformity as an explanation for force enhancement. For non-uniformity to be accepted, this concept must be extended to include halfsarcomere length non-uniformity (22). A-band displacements in preparations with myofibrils (38,39) and individual sarcomeres (37) were clearly detected during and after activation. There has been increasing evidence that residual force enhancement could be attributed to asymmetrical length changes inside one sarcomere that caused an increasing overlap in one of the two halves (21). A recent mathematical model (40) using a series of dissimilar halfsarcomeres (in several conditions) was able to generate a residual force enhancement on the order of 13% above the isometric reference contraction. Although exciting, this simulation was not able to produce higher levels of force enhancement or a duration longer than 30 s. According to the model, when both halves cease to lengthen or shorten, the different populations of cross-bridges reach an equilibrium, causing the force to decrease to the same level as the isometric reference. Hence, half-sarcomere length nonuniformity cannot fully account for the residual force enhancement. It is possible that simulations with more myofibrils in parallel could lead to a further increase in the magnitude and duration of the force enhancement found.

Hypothesis 2 -Engagement of a passive element
It has been suggested that force enhancement after stretch may also be related to an increase in strain in passive elastic elements inside the muscle (12). These elements may not, in fact, be totally ''passive;'' they become stiffer when muscles are stretched while activated. The most likely ''passive element'' for this function is titin (18).

Support for the theory
A series of studies (14,18,19,41) demonstrated an increase in the passive force even after the muscle was relaxed from the enhanced state ( Figure 1). This was called passive force enhancement. Generally speaking, this behavior is very similar to the actual total force enhancement; it is velocity independent, increases with stretch magnitude and is persistent (18,41,42). However, its magnitude is smaller than the total enhancement (18,42) and is thus assumed to account for only part of the phenomenon.
Previously, the origin of passive force enhancement was unknown, but it was commonly interpreted as the engagement of a passive element that becomes stiffer after stretch during activation and is not eliminated by deactivation (3,18). Passive and total force enhancements were shown to diminish when a shortening is applied prior to stretch, which was attributed to a release in the strain on elastic structures whose force was enhanced by the stretch (14). As mentioned previously, titin is considered the most likely candidate for this stretch-activation-sensitive role (16)(17)(18)43,44) because its stiffness can increase due to the effect of calcium binding to two specific motifs, the PEVK (Pproline, E-glutamine, V-valine, K-lysine), and immunoglobulin (Ig) domains, that have been found to change their molecular conformation upon calcium activation (45) and through binding to actin (46).
Increasingly, researchers have suggested that passive force enhancement may actually play a greater role than previously thought in contributing to the total residual force enhancement (18,42). It is believed that the greater the stretch magnitude, the greater the contribution of passive forces to the total force enhancement (18,42). Herzog and Leonard (18) found a greater than 80% passive force enhancement contribution to the total force enhancement at longer lengths. Even on the ascending limb of the forcelength relationship, where passive forces are not thought to play a significant role, a small passive force enhancement was detected (47). Indeed, Leonard and Herzog (44) recently showed that passive forces could be increased to almost 400% when relaxed myofibrils were compared with activated myofibrils while having their average sarcomerelength stretched from 2.4 mm to 6.0 mm. The authors attributed this notable increase in force to a combination of two mechanisms: 1) titin stiffening mediated by calcium activation and 2) a progressive binding of titin to actin through a mechanism that is likely to be force dependent and elicited by stretch after activation. This interaction between titin and actin was not a novel observation, but Leonard and Herzog (44) proposed a new understanding for its role in residual force enhancement. The authors also suggested that the high levels of force enhancement encountered could confer a protective mechanism against stretch-induced muscle injuries.

Limitations and criticisms
Passive forces can be modulated upon calcium activation (48), showing that an increase in stiffness can be crossbridge independent (49). As previously mentioned, this increase in stiffness is attributable to titin (18). Based on these results, it is assume that at least the passive force enhancement could be attributed to titin activation. However, titin stiffening via calcium binding only accounts for 25% of the total passive force enhancement (43). Therefore, calcium-induced titin stiffening alone is insufficient to explain even the passive part of force enhancement.
The explanation proposed by Leonard and Herzog (44) is attractive but has been criticized (50); although one study showed that titin can interact with actin (51), it remains unclear whether there is in fact interaction between PEVK and Ig and actin. It may be that stretches beyond the sarcomere length of 4.0 mm could have the effect of unfolding these domains and allowing their interaction with actin, but there is little evidence for this possibility (22,50).
The levels of passive force observed by Leonard and Herzog (44) do not agree with results from a similar study that also reported passive force modulated by a lengthdependent combination of stretch and activation (48). In this study, the greatest passive force increases found were approximately 30%. However, care should be taken when comparing results from these two studies because the latter used single fibers instead of myofibrils, and their average sarcomere length did not exceed 3.2 mm. Still, a 400% force increase is very unlikely since myofibrils are much more fragile than fibers, which also generate significantly lower levels of passive force (36,52). Furthermore, it is difficult to grasp why muscles would have such a strong protective mechanism, as Leonard and Herzog (44) suggest, if they are rarely stretched to that extent during muscle contractions in vivo.

Hypothesis 3 -Increase in the number of crossbridges
Another hypothesis to explain increasing tension in the enhanced state is an increase in the number of attached cross-bridges. There are several possible mechanisms at work: a stretch-induced phosphorylation of myosin light

Support for the theory
Several studies have found an increase in stiffness in the force-enhanced state compared with the isometric reference state in different muscle preparations (34,35,54). Assuming that most of the increase in muscle stiffness is due to an increase in the number of cross-bridges (55,56), these studies provide indirect evidence for our third hypothesis. For this increased stiffness to occur, according to Huxley's model (6), either the cross-bridge rate of detachment g (x) should decrease, or the rate of attachment f (x) should increase.
Stretch after activation may induce cross-bridge phosphorylation (57), myofilament-lattice disordering (53), and/ or repartitioning of the cross-bridge population between the ''force-bearing'' and ''non-force-bearing'' molecular states (58)(59)(60)(61). In the first case, the phosphorylated myosin light chains would bend myosin heads towards the actin filament (62,63). In the second case, myofilament disordering would cause the thin filaments to move closer to the thick filaments (53). In the third case, changes in cross-bridge distribution between the two aforementioned molecular states could induce an increase in the ''non-force-bearing'' state that would indirectly slow down the detachment rate after stretch because there would be more bridges in the ''nonforce-bearing'' state available to move forward to the ''forcebearing'' state (59,60). Finally, stretch per se could increase the time of attachment by slowing phosphate and ADP release, which has been previously observed in studies of single-molecule (myosin and actin) interactions (8,64,65).

Limitations and criticisms
The studies that compare stiffness in the force-enhanced state to the corresponding isometric state are few and contradictory. There are nearly as many studies that have shown either no change (24) or a decrease (29)(30)(31) in stiffness as studies that have shown increased stiffness (34,35,54). In addition, there are limitations in using stiffness measurements to make assumptions about the number of crossbridges, especially after stretch. Considering that this measurement would reflect the whole system's stiffness (i.e., titin, filaments, and the number of detached crossbridges in the ''non-force-bearing molecular state''), an increase in stiffness could also be attributed to an increase in strain on titin due to ''popped'' sarcomeres that would convey an enhanced stiffness from shorter, strong sarcomeres.
Even accepting that the number of cross-bridges is increased in the force-enhanced state due to a decrease in the myofilament space induced by stretch, which would in turn facilitate light-chain phosphorylation and force potentiation, as suggested by Edman (52), it is unlikely that stretch-induced myosin phosphorylation would be as long lasting as has been previously reported (1,3,4), which would be necessary for it to have a duration comparable to residual force enhancement. In fact, Edman (52) was concerned with characterizing force enhancement during force development; his method examined the force increase occurring right before the breakpoint on the force myogram, not after the force transients were absent. Therefore, the mechanism suggested by this author is plausible to explain these force transients, but it is unlikely to also explain residual behavior. Similarly, there is no reason to believe that stretch-induced myofilament disordering (53) or the redistribution of the cross-bridges between ''non-force-'' and ''force-bearing'' states (59,60) when considering that only the stretch effect could continue for several seconds after stretch, which is as long as the residual force enhancement has been reported (1,14,18,19). In other words, both mechanisms should be continually maintained for the cross-bridge-kinetics to remain permanently altered after the stretch, which is unlikely.
Finally, the results from studies of single myosin molecules with few myosin interactions that were observed for a brief time interval (64)(65)(66) are difficult to extrapolate to long-lasting phenomena such as residual force enhancement. This limitation is particularly true because residual force enhancement was studied in preparations at the single sarcomere level and up that involved several cross-bridge cycles occurring for many seconds after the stretch (1,4,10,13,15,(17)(18)(19).

Summary and Conclusions
Controversy remains concerning the three hypotheses discussed. Sarcomere length non-uniformity has been highly criticized not only because some of its predictions have been shown to be incorrect but particularly in light of recent results from muscle preparation studies that show force enhancement even at the single sarcomere level. For non-uniformity to be considered the exclusive mechanism of residual force enhancement, the concept of half-sarcomere length non-uniformities causing force increase must be more thoroughly tested experimentally.
There is sufficient evidence showing that a passive elastic element (i.e., titin) could also be responsible for force enhancement at longer lengths. However, it is unlikely that this is the exclusive mechanism since the passive force enhancement is always smaller than the total force. In fact, the cause of passive force enhancement is still not entirely clear; experiments that indirectly evaluated titin activation mediated by stretch after activation were able to attribute only 25% of the passive force enhancement to titin activation. Furthermore, findings on passive force regulation are contradictory. Similar studies have obtained very different values for increased force magnitude.
There is some evidence showing an increase in the number of cross-bridges when comparing the forceenhanced with the corresponding isometric state. It appears that a stretch after activation can produce long-lasting changes in cross-bridge kinetics. However, the exact mechanism for this remains unclear.
Thus far, none of the three hypotheses provide a full explanation, although none can be completely ruled out. Evidence has shown that sarcomere length non-uniformity alone can no longer explain residual force enhancement. To do so, the concept must be extended to include halfsarcomere length non-uniformity as a source of force enhancement, although it is also unlikely half-sarcomere length non-uniformity alone could explain force enhancement. Therefore, based on several strong but partial pieces of evidence and the contradictions from the three hypotheses presented here, we suggest that residual force enhancement is perhaps caused by all three mechanisms operating together: i) engagement of a passive element, likely via titin activation and interaction with actin, ii) dissimilarities in half-sarcomere lengths, and iii) an increase in the number of cross-bridges due to alterations of their kinetics caused by stretch after activation.

& AUTHOR CONTRIBUTIONS
Minozzo FC and Lira CA contributed to manuscript preparation, interpretation, and critical revision of the manuscript.