Fecal incontinence affects people of all ages and social backgrounds and can have devastating psychological and economic consequences. This disorder is largely attributed to decreased mechanical efficiency of the internal anal sphincter (IAS), yet little is known about the pathophysiological mechanisms responsible for the malfunction of sphincteric smooth muscle at the cellular level. The object of this study was to develop a three-dimensional (3-D) physiological model of the IAS bioengineered in vitro from isolated smooth muscle cells. Smooth muscle cells isolated from the IAS of rabbits were seeded in culture on top of a loose fibrin gel, where they migrated and self-assembled in circumferential alignment. As the cells proliferated, the fibrin gel contracted around a 5-mm-diameter SYLGARD mold, resulting in a 3-D cylindrical ring of sphincteric tissue. We found that 1) the bioengineered IAS rings generated a spontaneous basal tone, 2) stimulation with 8-bromo-cAMP (8-Br-cAMP) caused a sustained decrease in the basal tone (relaxation) that was calcium-independent, 3) upon stimulation with ACh, bioengineered IAS rings showed a calcium- and concentration-dependent peak contraction at 30 s that was sustained for 4 min, 4) addition of 8-Br-cAMP induced rapid relaxation of ACh-induced contraction and force generation of IAS rings, and 5) bioengineered sphincter rings show striking functional differences when compared with bioengineered rings made from isolated colonic smooth muscle cells. This is the first report of a 3-D in vitro model of a gastrointestinal smooth muscle IAS. Bioengineered IAS rings demonstrate physiological functionality and may be used in the elucidation of the mechanisms causing sphincter malfunction.
- extracellular matrix
- 8-bromo-adenosine 3′,5′-cyclic monophosphate
fecal incontinence is associated with reduced anal closure pressure and mechanical stress for maintaining tissue coaptation and closure of the anal canal, resulting in the inability to control the passage of gas or stools (feces) through the anus (11, 49). This is largely attributed to decreased mechanical efficiency of the internal anal sphincter (IAS; see Refs. 28, 32, and 48). Epidemiological studies have shown that fecal incontinence affects 2–15% of the population (15, 41, 44, 50), including people of all ages and social backgrounds, thus making it one of the most common gastrointestinal disorders. A diagnosis of fecal incontinence often leads to devastating social, psychological, and economic consequences and a significantly lower quality of life (44). Despite the prevalence and disturbing nature of this condition, little is known about the mechanisms underlying smooth muscle sphincter malfunction. Thus there is a need for a model that provides a comprehensive understanding of sphincteric smooth muscle physiology.
From a clinical perspective, understanding sphincter malfunction in regard to fecal incontinence has been limited because traditional anal manometric measures are not able to localize specific damage or deterioration to individual sphincter structures or discern the underlying cellular or tissue processes responsible for function loss. Studies have tended to focus on symptomatic individuals and on structure or function, but not both. Furthermore, without a comprehensive understanding of the pathophysiological mechanisms responsible for sphincter malfunction at the cellular level, the development of corrective strategies and treatment options for people diagnosed with fecal incontinence have been limited.
Tissue culture was developed in the early 1900s as a technique for studying the behavior of animal cells in vitro. Advances in technology have allowed the use of tissue culture for many areas of study, including intracellular activity, intracellular flux, environmental interaction, cell-cell interaction, and genetics (17). The capacity to grow different cell types in culture and subsequently test the effects of a variety of drugs, hormones, and other interventions on cellular function is of the utmost importance. However, when force production is the primary tissue-level function, two-dimensional cell culture is an inadequate model for most experimental needs. Furthermore, cells that are propagated and tested two-dimensionally lack many of the cellular and environmental cues that play a role in differentiation and the establishment of normal physiological function. Isolation of single cells and cell suspensions has enabled biochemical and mechanical measurements at the cellular level and has advanced our level of understanding of smooth muscle function (6, 36). Yet cell suspensions also lack the potential for cell-cell interactions and cell-matrix interactions provided by three-dimensional (3-D) tissues (17) and lack the tissue-level organization and system-level function to allow a direct study of the underlying mechanisms of clinical importance (14).
It is well known that physiological functions of tissues are retained when their 3-D structure is kept intact (22). Isolated tissues and organ preparations, such as muscle strips, have been shown to provide researchers with a 3-D tissue that can be easily subjected to controlled changes in perfusion, oxygen availability, and agonist-induced stimulation (19, 20, 31). Despite the obvious advantages of tissue/organ explants, many limitations remain. Tissues are composed of several different cell types. For example, IAS muscle strips may be comprised of smooth muscle but also any combination of mucosal, epithelial, and/or neuronal cells. Thus it is difficult to experimentally manipulate or investigate one tissue constituent. Explanted tissues do not last indefinitely and are usually only viable for up to 4 h after removal, which prevents long-term investigation. Because the tissues must be manipulated for experimental purposes almost immediately after dissection, they must be prepared de novo for each experiment. Damage to the tissue and/or release of material from damaged erythrocytes occurring during dissection also inhibits the ability to produce normal functionality and environmental conditions. Last, tissue/organ explants must be placed in a cold (usually 4°C) buffer or bath to prevent rapid degradation. For these reasons, tissue engineering has emerged as a valuable tool that applies the principles of engineering and life sciences toward the development of biological models with characteristics similar to those observed in vivo. Specific cell types can be isolated and bioengineered to yield a homogenous tissue. Bioengineered tissues can be maintained in culture for long periods of time under physiological conditions (12–14). Advances in tissue engineering have been clinically applied to restore, maintain, and improve tissue function, resulting in enhanced treatment for damaged tissues (16).
There have been many reports of 3-D “tissue equivalents” developed in vitro by culturing cells in special matrixes or under conditions to promote self-organization (13, 14, 23, 46). Fibrin-based gels have become an effective option for engineering functional striated muscle tissue, providing a matrix that is well suited for the development of contractile tissues (39, 46). Fibrin gels are formed by the enzymatic cleavage of fibrinogen by the serine proteinase thrombin, allowing the fibrin monomers to spontaneously interact and form fibrils. Fibrin-based gels have advantages over synthetic polymer approaches of tissue-equivalent fabrication, specifically immediate cellularity of the construct through cell entrapment in the forming gel and subsequent fibril alignment via mechanical constraint of cell-induced gel compaction (46). Cell-mediated fibrin gel contraction occurs, mimicking alignment of muscle tissue in vivo (39). Within a fibrin matrix, cells rapidly migrate, proliferate, and digest the fibrin, replacing it with their endogenous extracellular matrix (ECM; see Refs. 21, 22, 40, and 46). Vascular smooth muscle cells grown in fibrin have shown increased production of ECM, collagen, (22, 40), and elastic fibers (34) and also exhibited mechanical properties that are commensurate with vascular tissue in vivo (34, 46). Furthermore, fibrin-based engineered tissues can be maintained for weeks in culture (23, 46). The prolonged life span of engineered tissues allows temporal investigation of the effects of cell-cell interactions, growth factors, mechanical and chemical stimuli, gene expression, and kinase action on tissue-level function.
Here we used fibrin gel casting to develop an in vitro model of the IAS from isolated rabbit smooth muscle cells. Our bioengineered “IAS rings” are 3-D cylindrical structures resembling the IAS portion of the gastrointestinal tract. Isolated IAS cells seeded in a fibrin construct self align and form a ring in ∼5–10 days. The resulting IAS rings are functionally similar to gastrointestinal smooth muscle in vivo for >3 wk, including generation of spontaneous tone and agonist-induced contraction and relaxation of the basal tone with the appropriate stimuli. Stimulation with 10−5 M 8-bromo-cAMP (8-Br-cAMP) resulted in a calcium-independent relaxation of −16.3 ± 2.0 μN from the basal tone. When stimulated with ACh, the IAS rings produced a calcium-dependent maximal force of 15.7 ± 1.4 μN. ACh-induced stimulation was also concentration dependent, with a plateau at 10−6 M. Addition of 8-Br-cAMP induced rapid relaxation of ACh-induced contracted rings. Last, isolated colonic smooth muscle cells were bioengineered into 3-D rings and tested in the same manner as sphincteric rings. Bioengineered colon rings exhibited spontaneous phasic contractile behavior (amplitude: 18 ± 1.99 μN; frequency: 2.45 ± 0.21 cycles/min). Sphincter rings showed marked functional differences from colonic smooth muscle rings, including the absence of spontaneous phasic contractile behavior. This is the first report of a 3-D cylindrical, in vitro model of the IAS that is functionally similar to smooth muscle in vivo. This model provides the opportunity for a novel approach to investigate smooth muscle function and dysfunction, including the elucidation of disease mechanisms causing fecal incontinence.
MATERIALS AND METHODS
Materials and solutions.
Growth media (GM) consisted of DMEM (catalog no. 12430–054; GIBCO) with 15% FBS, 3% penicillin-streptomycin, and 0.6% l-glutamine. Differentiation media (DM) consisted of 73% DMEM (catalog no. 12430–054; GIBCO), 20% Media 199, 7% heat-inactivated horse serum, and 1% penicillin-streptomycin. Krebs solution contained (in mM) 119 NaCl, 4.6 KCl, 15 NaHCO3, 1.5 CaCl2, 1.2 MgCl2, 1.2 NaHCO3, and 11 glucose; 0 Ca2+/2 mM EGTA Krebs solution was the same except CaCl2 was not included and 2 mM EGTA was added. HEPES buffer (pH 7.4) contained (in mM) 115 NaCl, 5.7 KCl, 2.0 KH2PO4, 24.6 HEPES, 1.9 CaCl2, 0.6 MgCl2, 5.6 glucose, 0.01% soybean trypsin inhibitor, and 0.184 (wt/vol) DMEM; 0 Ca2+/2 mM EGTA HEPES solution was the same except CaCl2 was not included and 2 mM EGTA was added.
Smooth muscle cells were isolated from the IAS of New Zealand White rabbits as described previously (4, 5). Briefly, the IAS (consisting of the most distal 3 mm of the circular muscle layer, ending at the junction of skin and mucosa) and the distal colon were removed by sharp dissection. The tissues were rapidly cleaned and stripped of connective tissue in ice-cold carbonated Krebs solution containing 2% penicillin-streptomycin. Separately, the tissues were cut into small pieces and transferred to a 100-mm plate containing 15 ml HEPES buffer with 0.1% collagenase type II (Worthington Biochemical) for digestion. The plates were placed in an incubator (37°C with 5% CO2) for a 1-h digestion period. Fresh HEPES (15 ml) buffer with collagenase was added, and the contents of the plates were mechanically dissociated using a 10-ml pipette and incubated for one additional hour. After the second digestion period was complete, the cells were collected and centrifuged at 800 g for 10 min, and the supernatant was discarded. The cells were washed and resuspended three times to ensure removal of excess collagenase and then resuspended in HEPES buffer or 0 Ca2+/2 mM EGTA HEPES buffer.
ACh-induced stimulation and measurement of contraction in suspension of isolated IAS smooth muscle cells.
To determine the effect of ACh on contraction of isolated sphincteric smooth muscle cells, ACh was added to 0.5 ml suspension of isolated smooth muscle cells for 30 s or 4 min, with untreated cells as controls. The reaction was stopped, and cells were fixed by the addition of 0.1 ml acrolein at a final concentration of 0.1%. Individual cell lengths were measured by computerized image micrometry (5, 7). The length of cells in the control state or after addition of ACh was obtained by measuring 30–50 cells encountered randomly in successive microscopic fields from each of three separate experiments. The contractile response is defined as the decrease in the average length of the cells counted and is expressed as the percent change from the control length.
Preparation of culture dishes and culture of rings.
Culture plates (35 mm) were prepared as described previously (12, 13). Briefly, 1.5 ml SYLGARD [polydimethylsiloxane (PDMS)] was poured in each plate and allowed to cure for 24 h. The SYLGARD substrate provided a layer in which anchor materials could be easily pinned in place and provided a surface to which, unless coated, the cells could not adhere. A cylindrical SYLGARD mold with an upwardly protruding 5-mm-diameter disk was placed in the center of the dish to provide a luminal space for the engineered sphincter or colon (Fig. 1A). Adhesion of the mold to the SYLGARD-coated culture dish was facilitated by pressing the mold firmly against the bottom of the dish with forceps. The culture dish was sterilized with 70% ethanol for 30 min. Ethanol was then aspirated, and culture dishes were exposed to ultraviolet light for 1 h. Each dish received 500 μl GM containing 10 U/ml thrombin, and the dish was agitated until the bottom of the plate was entirely coated. Finally, 200 μl of 20 mg/ml fibrinogen was added to each plate and gently swirled. The fibrin polymerized ∼15 min later, and the plates were ready for cell seeding.
Primary cells isolated from rabbit IAS and colon tissues were expanded separately in 75-cm2 flasks for ∼1 wk. The cells were then detached from their culture flasks with 1.5 ml trypsin-EDTA, collected, resuspended in GM, and counted with a hemocytometer. The cell density was adjusted, and 5 × 104, 7.5 × 104, or 10 × 104 cells/ml was added to the plates in a final volume of 2 ml. Plates were returned to the incubator and removed every 48 h to change the media. After plating (5–10 days, depending on the density of the cells), GM was replaced with DM to promote the formation of adult cellular connections and smooth muscle differentiation (23, 46). The DM was replaced every 2–3 days until the ring was fully contracted and ready for testing. IAS and colon rings were fully formed in ∼5–10 days; however, time spent in culture for each construct varied depending on the number of cells plated and the experimental design designated for each construct.
In parallel, fibrin gel constructs were prepared normally; however, cells were not added before the incubation period to characterize the mechanical properties of the fibrin gel in the absence of cells.
Measurement of contractile properties.
The protocol for measuring excitability and contractility of engineered muscle constructs was adapted from previous work (12–14, 23). The following three variables were directly measured: torroidal diameter of the muscle construct, passive baseline force (Pb), and peak change in isometric force (ΔP). The following two additional parameters were then calculated from the measured variables: cross-sectional area (CSA) was calculated from the diameter and specific force (sΔP) by dividing ΔP by the CSA. Briefly, engineered sphincters and colons were separated from their molds (Fig. 1E) using forceps, and the minimum ring diameter was measured using a calibrated eyepiece and a 5 or 10× objective lens on an inverted microscope (Axiovert 25; Zeiss, Thornwood, NY). The cross section was circular; therefore, the CSA was calculated using the measured diameter.
DM in the plate was replaced with 37°C Krebs solution, and the dish was placed on a heated aluminum platform that was maintained at a temperature of 37°C until the testing was complete. For contractility measurements, one end of the engineered sphincter or colon was anchored by a stainless steel pin (10 mm × 0.1 mm diameter) to the PDMS substrate, whereas another stainless steel pin was bent in the shape of a hook and attached by canning wax to an optical force transducer with a resolution of 1.4 μN and a range of 2 mN (Fig. 1, F and G; see Refs. 12, 13, 35, and 37). We followed standard protocols used for measuring spontaneous basal tone of IAS muscle strips as follows: stretching of the tissue, followed by a period of equilibrium (between 20 and 60 min), where the IAS strips stabilized, resulting in the establishment of a new stable baseline of tension (2, 19, 20, 31). Thus the bioengineered IAS and colon rings were stretched ∼50% of their resting length using a three-axis micromanipulator. The rings were allowed to sit for 20–30 min to reestablish a stable Pb, which was arbitrarily set at zero to allow consistent measurements of ΔP.
Agonist-induced stimulation of bioengineered rings.
All force measurements were collected at 100 samples/s for 60 s and recorded using a computer with LabVIEW data acquisition software (National Instruments, Austin, TX). A median filter of rank 2 was applied to all raw force data before being stored to minimize digitization noise without causing a phase delay for rapidly changing forces. ΔP was determined by subtracting the Pb from the total force values. Data analysis was done using LabVIEW and Microsoft Excel software programs. Each measurement was repeated three to eight times, and the mean value was recorded.
Contractions were induced in the rings by the addition of 10−10 to 10−6 M ACh. Maximal contractions were seen ∼30–60 s after addition of the drug. Relaxation response in the rings was induced by the addition of 10−5 M 8-Br-cAMP. Maximal relaxation was achieved after ∼4 min; therefore, relaxation measurements are collective of four consecutive 1-min data recordings.
Preparation for histology.
In preparation for light microscopy, engineered IAS rings were fixed in 4% paraformaldehyde. The rings were processed and embedded in paraffin wax according to standard histological procedures (9). All sections were made (5–8 μm thick) using a rotary microtome. Staining of the histological sections was performed to view general structure using Harris's hematoxylin and eosin (24).
Tissue engineering data are presented as means ± SE for three to eight experiments per data point. Differences in mean values were compared within groups (e.g., control vs. ACh treatment), and significant differences were determined by ANOVA with the post hoc Tukey-Kramer Honest Significant Difference test. The level of significance was set at P < 0.05.
In designing an innovative model of the IAS, the following parameters were considered: three dimensionality, remodeling of in vivo structure, expression of fundamental in vivo physiological behaviors, potential for cell-cell and cell-matrix interactions, enhanced viability, and maintenance under physiological conditions in a controlled environment.
Contractility of isolated IAS cells.
ACh (10−7 M)-induced stimulation of isolated smooth muscle cells from the rabbit IAS resulted in a sustained contractile response. Contraction was defined as the percent decrease in cell length compared with control cell length (86.86 ± 5.29 μm, n = 150). ACh (10−7 M) induced a peak contraction at 30 s (28.85 ± 3.20% decrease in cell length, n = 150) and was sustained for 4 min (27.09 ± 4.43% decrease in cell length, n = 150; Fig. 2). To determine the calcium dependence of this response, parallel experiments were performed in 0 Ca2+/2 mM EGTA HEPES buffer. Control length of isolated smooth muscle cells preincubated in 0 Ca2+/2 mM EGTA was 88.68 ± 6.41 μm (n = 150). As expected, ACh-induced stimulation of isolated smooth muscle cells resulted in minimal contractile response in the absence of Ca2+ (0.5 ± 4.47 and 4 ± 3.91% decrease in cell length at 30 s and 4 min, respectively; Fig. 2). These data are similar to the previously published contractile response of isolated IAS smooth muscle cells to bombesin and to exogenous protein kinase C (3, 4).
Bioengineering a 3-D IAS or colon ring.
Primary cells isolated from the IAS and the circular smooth muscles of the colon were cultured separately in 75-cm2 flasks. After ∼1 wk, 50,000 or 100,000 cells were transferred to plates containing a loose fibrin gel where they proliferated to confluence. As expected, cells proliferated and began to digest the fibrin gel, replacing it with endogenous ECM (21–23, 40, 46). Cells migrated and self-assembled along the line of force until they formed a parallel array of cells. As cells proliferated, they shrunk the fibrin gel, contracting it toward the center of the culture dish around a 5-mm-diameter SYLGARD mold (Fig. 1, B-D). The rate of gel contraction was dependent on the number of cells initially plated, since constructs seeded with 100,000 cells consistently formed rings faster than constructs seeded with 50,000 cells (data not shown). Tissues were considered “formed” when a 3-D cylindrical tube of sphincteric or colonic tissue contracted around the SYLGARD mold (Fig. 1D). All fibrin constructs seeded with IAS or colon cells were fully formed within 5–10 days after cell seeding. The resulting 3-D IAS and colon rings remained stable in culture and were experimentally tested between 9 and 23 days.
Parallel constructs of gels made without the addition of cells did not exhibit any contraction of the gel, and the gels without cells appeared unchanged after 21 days in culture. Thus contraction of the cell around the SYLGARD mold is mediated by the cells seeded in the construct.
Histological analysis of the bioengineered IAS rings revealed one uniform cell type surrounded by undigested pieces of the fibrin gel matrix (Fig. 3). The cells had many features of differentiated smooth muscle cells (fusiform, nonstriated, and uninucleated with the nucleus in the center of the cell), although the volume of the perinuclear space was small compared with cells in a normal muscle tissue. This suggests that the cells had not reached full maturation. As the gel contracted over a period of days, cells increasingly aligned in a parallel array. Histological analysis showed a gradient of cell alignment as the gel contracted and formed a bioengineered IAS ring (Fig. 3). Fully contracted rings were composed of ∼20 concentric cell layers. Ring diameters averaged 91.3 ± 13.6 μm and had a range of 46.5–145.4 μm. The average CSA was calculated as 7,532.9 ± 2,010.3 μm2 and ranged from 1,697.40 to 16,604.90 μm2. These measurements were used to determine the specific force of each IAS ring, defined as the maximal force produced divided by the CSA.
Development and relaxation of the basal tone in bioengineered IAS rings.
Elongation of an IAS ring by ∼50% of its length resulted in an elevation of tension followed by relaxation that was initially substantial, but gradually subsided over a period of 20–30 min as a stable baseline of tension was reached. This stable baseline was arbitrarily set at zero.
Upon the addition of 10−5 M 8-Br-cAMP, IAS rings demonstrated a sustained relaxation response (reduction of the stable basal tone) reaching maximal relaxation ∼4 min after stimulation and subsequently resulting in the reestablishment of a new, stable basal tone. Average ΔP for the rings after 4 min of incubation with 10−5 M 8-Br-cAMP was −14.8 ± 2.56 μN (Fig. 4B). Stimulation with 10−5 M 8-Br-cAMP in 0 Ca2+/2 mM EGTA Krebs also resulted in a relaxation response (−23 ± 4 μN after 4 min), suggesting that relaxation was calcium independent (Fig. 4).
ACh-induced contraction of bioengineered IAS rings.
Bioengineered IAS rings stimulated with 10−6 M ACh exhibited a peak ΔP of 10.34 ± 3.2 μN within 30–60 s and was sustained for 4 min (Fig. 5). Maximal ΔP induced by ACh stimulation was 33.7 μN, resulting in a specific force (sΔP) of 2.8 kN/μm2. Rings in 0 Ca2+/2 mM EGTA Krebs did not contract when stimulated with ACh (0.9 ± 0.2 μN; Fig. 5), demonstrating the calcium dependence of the contractile response. Furthermore, force generated by ACh-induced stimulation was concentration dependent (Fig. 6), reaching a plateau at 10−6 M.
ACh-induced contraction of IAS rings and subsequent relaxation with 8-Br-cAMP.
ACh-induced contraction of IAS rings did not appear to effect subsequent relaxation induced by 10−5 M 8-Br-cAMP. Addition of 10−5 M 8-Br-cAMP resulted in an immediate relaxation of ACh-induced contracted IAS rings (Fig. 7). Addition of 8-Br-cAMP induced rapid relaxation of ACh-induced contraction and force generation of IAS rings (−14.3 ± 3.8 μN; Fig. 7B).
Spontaneous phasic contractile behavior in bioengineered colon rings.
ACh-induced contraction and 8-Br-cAMP-induced relaxation of bioengineered colon rings showed a similar response pattern to that seen in bioengineered sphincteric rings (data not shown). However, colon rings showed one striking functional difference from IAS rings; they exhibited spontaneous phasic contractile behavior (Fig. 8). Without the addition of any external stimuli, 100% of bioengineered colon rings demonstrated spontaneous phasic contraction. Spontaneous phasic contractions were recorded before, during, and after experimental testing (while the ring was still attached to the recording equipment), depending on the individual ring. Data were recorded at 1-min intervals continuously until contractions stopped. If uninterrupted, spontaneous phasic contractions persisted (amplitude: 18 ± 1.99 μN; frequency: 2.45 ± 0.21 cycles/min), on average, for 10.3 ± 2.19 min. Spontaneous phasic contractions were not seen in any IAS constructs at any time during data recording.
Despite the prevalence of fecal incontinence, little is known about the mechanisms underlying sphincter malfunction. Previous research attempts to understand IAS physiology, including tissue culture (17), cell suspensions (1, 3, 4), and tissue explants (1, 19, 20, 31), have advanced our knowledge of smooth muscle function. However, each technique has its own limitations; thus, smooth muscle function is still not clearly understood. New technological advances in tissue engineering, cell survival, and the utilization of 3-D matrixes offered new opportunities to engineer 3-D IAS sphincters. Tissue engineering incorporates three dimensionality resembling tissue explants and a homogenous population of cells similar to cell suspensions and utilizes tissue culture techniques to provide a controlled environment under physiological conditions in which tissues can survive for extended periods of time. More recently, fibrin-based tissue engineering of arterial smooth muscle has revealed a promising new technique for investigating smooth muscle (46). Using fibrin gel casting, we have developed a 3-D model of the IAS in vitro from isolated rabbit smooth muscle cells. This system uses a homogenous population of isolated sphincteric smooth muscle cells and, when seeded on a fibrin gel, they self-organize, forming 3-D functional tissues. In this study, all IAS rings formed in ∼5–10 days. Previous work has shown fibrin-based tissue construct viability for up to 40 days (23). Our IAS rings were only tested between 9 and 23 days in culture and therefore provide evidence for viability beyond 3 wk. Bioengineered IAS rings demonstrate spontaneous tone (force produced) that decreased (relaxation) in a calcium-independent manner by the addition of 8-Br-cAMP or increased (contraction) in a calcium-dependent manner by the addition of ACh. Addition of 8-Br-cAMP also induced rapid relaxation of ACh-induced contracted IAS rings (Fig. 7). These data provide evidence of physiological functionality. In addition, the absence of spontaneous phasic contractile behavior in IAS rings, which was seen in 100% of bioengineered colon rings, demonstrates tissue specificity. This model is highly reproducible and will offer an effective method for investigation of smooth muscle function because it combines the major advantages of previously reported research methods into one technique.
Bioengineered IAS rings exhibited physiological behavior functionally similar to smooth muscle in vitro. Upon seeding, cells began digesting the fibrin matrix, replacing it with their own ECM (21–23, 40, 46). As the cells proliferated, they self-organized along the line of force, forming a parallel array of cells analogous to what is observed in 3-D and mechanically strained cultures (29, 30, 33, 47). The rate of ring formation was dependent on the number of cells initially plated, although this did not appear to affect force generated, since the magnitude of force generated by IAS rings was consistent regardless of the seeding number (data not shown). This was consistent with previous studies involving fibrin-based muscle constructs (23). The cells appeared to proliferate within the fibrin gel only until they reached confluence, regardless of seeding density. This suggested that normal density-dependent contact inhibition of cell division (8) had occurred during the formation of IAS rings.
Agonist-induced stimulation showed that bioengineered rings were functionally similar to smooth muscle in vivo. IAS rings demonstrated a stable, spontaneous tone until stimulated by an agonist. IAS rings showed sustained calcium-independent relaxation of the basal tone in response to 10−5 M 8-Br-cAMP. This is similar to smooth muscle in vivo, where relaxation is largely mediated by cAMP and is calcium independent (10, 27, 38, 39). Upon stimulation with the same concentration of ACh, both IAS rings and isolated IAS single cells exhibited a peak ΔP within 30–60 s after stimulation that was sustained for 4 min (Figs. 2 and 5A). Contraction of both isolated single cells and bioengineered rings were calcium dependent (Figs. 2 and 5A). These similarities suggest that bioengineered IAS rings have the sensitivity of isolated single cells, which may be because of the lack the barriers present in muscle strips, such as innervating nerves, epithelial lining, and serosa, and therefore provide an in vitro opportunity to experimentally manipulate one 3-D tissue constituent. ACh-induced contraction of both isolated single cells and bioengineered rings also corresponds to previously described ACh-induced contraction of human IAS muscle strips (19, 20, 31). Preincubation of engineered IAS rings with 10−6 M ACh did not affect the relaxation response induced subsequently with 10−5 M 8-Br-cAMP, since IAS rings demonstrated an immediate shift from contraction to relaxation after 8-Br-cAMP-induced stimulation (Fig. 7). Bioengineered IAS rings have the ability to contract and relax within minutes, similar to the capacity seen in vivo. Therefore, 3-D engineered IAS constructs exhibited several physiologically functional behaviors similar to the capacity seen in vivo.
The characteristic formation of bioengineered IAS rings from isolated sphincteric cells is exclusive to the IAS. Bioengineered colon constructs demonstrated spontaneous phasic contraction in the absence of external stimuli. Bioengineered colon constructs were made from isolated colonic cells from the same animals used to make IAS rings. The frequency of bioengineered colonic phasic contractions (2.45 ± 0.21 cycles/min) was similar in frequency to that seen in colonic muscle strips (2–4 cycles/min; see Ref. 45). As expected, bioengineered IAS rings did not exhibit the spontaneous phasic contractile behavior at any time. This suggests that IAS constructs, prepared using adjacent portions of the gastrointestinal tract from the same animals, exhibited physiological functionality and characteristic tissue specificity that are unique to the IAS.
Although the engineered rings described here provide an important model for studying sphincteric smooth muscle, it is important to note that these cells were not fully mature. Histological analysis of the bioengineered rings revealed a homogenous composition of differentiated smooth muscle cells that had not reached full maturation. Maturation of smooth muscle cells involves continuous regulation by a variety of environmental factors, including hormonal signals and neural and mechanical input (25, 43). Although it is known that these factors influence the maturation of smooth muscle cells, the specific role of each factor in vivo is not clearly understood (43). Optimization experiments for IAS constructs could be designed in which each physiological input is altered to maximize cell maturation. This would improve the engineered IAS rings but would also independently provide a better understanding of smooth muscle maturation in general. Ross and Tranquillo (46) revealed increased growth of smooth muscle cells in a fibrin matrix (measured by collagen production) associated with GM (containing DMEM supplemented with FBS) vs. DM (consisting mainly of media 199). After formation of the ring, we attempted to promote differentiation by replacing the normal GM with a differentiated GM that has fewer mitogenic signals, therefore suppressing mitosis. This demonstrates the association between maturation and increased force production. Although our IAS rings were developmentally arrested and had not reached full maturation, this model is well suited for investigation of smooth muscle development, maturation, and maintenance of functional smooth muscle.
Although fibrin gel-based constructs have previously been studied, a quantitative comparison of force generation was not possible. Tranquillo and colleagues (18, 21, 26, 40, 46) provide the necessary foundation for this area of research, demonstrating and characterizing arterial smooth muscle tissue growth and development within a fibrin matrix and the resulting tensile properties. However, fibrin-based engineering of smooth muscle has been limited to vascular/arterial smooth muscle. Furthermore, there are no data available that report the active forces generated by fibrin-based smooth muscle constructs. This is the first report of active forces, including specific force, generated by fibrin-based smooth muscle constructs. Furthermore, this paper provides the first illustration of not only gastrointestinal smooth muscle, but sphincteric smooth muscle bioengineered in a fibrin matrix. Our research on bioengineered IAS rings advances fibrin-based engineering of smooth muscle tissue by investigating and correlating the functional properties, including measurements of ΔP in response to agonist-induced stimulation. Specific force calculated for bioengineered IAS rings was 2.8 kN/m2. The only available comparison of force generated by smooth muscle was explanted muscle strips from humans (10.2 kN/m2; see Ref. 19), which is 3.6-fold more force than the IAS rings. Nerem and Ensley (42) report significant variation in the mechanical strength of smooth muscle grown in artificial scaffolds among different species. Therefore, the lower force generation induced by ACh in these engineered IAS rings could be in part attributed to the different model systems being compared (human muscle vs. rabbit muscle). The difference in specific force generated in human muscle strips vs. bioengineered IAS rings may also be the result of the developmental state of the cells, since our system has not yet incorporated all of the environmental cues necessary for smooth muscle to reach full maturation. Furthermore, bioengineered rings were made using a homogenous smooth muscle population grown in a fibrin matrix. Although it has been shown that cells grown in a fibrin matrix rapidly migrate, proliferate, and then digest the fibrin, replacing it with their own ECM (21, 22, 40, 46), fibrin itself is compliant compared with the rigid ECM seen in vivo and therefore will not transmit force as well. Histology revealed that the bioengineered rings were composed of cells surrounded by pieces of undigested fibrin matrix. To address this issue in future experimentation, growth factors could be added to increase ECM production. For example, transforming growth factor (TGF)-β has previously been shown to increase the amount of ECM in fibrin-based engineered tissues (46). Incorporating TGF-β into our system may therefore increase ECM production and cell differentiation, resulting in a less compliant ring with enhanced force production and transmission. Furthermore, alteration of the method in which the cells are seeded may also lead to increased force generation in bioengineered constructs, for example, mixing the isolated cells directly with the fibrin before polymerization, rather than after, to increase homogeneity of the ring and reduce the percentage of undigested fibrin comprising the ring. Future research will be required to optimize this system using improved contractility as the outcome measure.
In conclusion, a fibrin-based model of the IAS has been produced. In this model, IAS cells self-organize in a 3-D fibrin matrix. The resulting 3-D rings show tissue specificity to the IAS, are highly reproducible, and are functionally similar to IAS smooth muscle in vivo. Our fibrin-based constructs provide the opportunity to test the effects of various pharmacological agents, growth factors, and mechanical interventions on smooth muscle function. This is the first report of a functional in vitro model of the IAS that may be used in the elucidation of the mechanisms causing smooth myogenic sphincter malfunction and the investigation of treatments for fecal incontinence.
This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant 5 RO1 DK-042876.
We thank the members of the Bitar laboratory and Richard Dapson for technical assistance. We thank Kelly Smid and Shilow Blea for assistance with technical editing and figure preparation.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
- Copyright © 2005 the American Physiological Society