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Am J Physiol Cell Physiol 291: C718-C725, 2006. First published August 23, 2006; doi:10.1152/ajpcell.00127.2005
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PROTEIN AND VESICLE TRAFFICKING, CYTOSKELETON

Chondrocyte intracellular calcium, cytoskeletal organization, and gene expression responses to dynamic osmotic loading

Pen-hsiu Grace Chao,1,2 Alan C. West,2 and Clark T. Hung1

1Department of Biomedical Engineering and 2Department of Chemical Engineering, Columbia University, New York, New York

Submitted 17 March 2005 ; accepted in final form 24 April 2006


   ABSTRACT
TOP
 ABSTRACT
METHODS AND MATERIALS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
While chondrocytes in articular cartilage experience dynamic stimuli from joint loading activities, few studies have examined the effects of dynamic osmotic loading on their signaling and biosynthetic activities. We hypothesize that dynamic osmotic loading modulates chondrocyte signaling and gene expression differently than static osmotic loading. With the use of a novel microfluidic device developed in our laboratory, dynamic hypotonic loading (–200 mosM) was applied up to 0.1 Hz and chondrocyte calcium signaling, cytoskeleton organization, and gene expression responses were examined. Chondrocytes exhibited decreasing volume and calcium responses with increasing loading frequency. Phalloidin staining showed osmotic loading-induced changes to the actin cytoskeleton in chondrocytes. Real-time PCR analysis revealed a stimulatory effect of dynamic osmotic loading compared with static osmotic loading. These studies illustrate the utility of the microfluidic device in cell signaling investigations, and their potential role in helping to elucidate mechanisms that mediate chondrocyte mechanotransduction to dynamic stimuli.

cartilage; calcium signaling; actin cytoskeleton; aggrecan

ARTICULAR CARTILAGE serves as the load-bearing material of joints, with favorable friction, lubrication, and wear characteristics (11, 41). The cells in cartilage, chondrocytes, are embedded in a matrix of white and dense connective tissue consisting of two phases, a solid organic matrix (50% collagen fibrils and 20–30% proteoglycan molecules) and a mobile fluid phase (predominately water) (15, 34, 43). Aggregates of proteoglycans are important in the development and maintenance of cartilage. Chondrocyte gene expression of aggrecan, a major component of these aggregates, can be modulated by chemical and mechanical forces (19, 49, 57). Because of its polyanionic glycosaminoglycan chains, the proteoglycan draws water into the tissue, resulting in an internal swelling pressure (6, 26, 32, 34, 42). The osmotic conditions of articular cartilage in situ have not been measured and have been estimated only empirically from phenomenological models (30, 31, 56) or theoretical multiphasic models (32). Degradation of matrix molecules occurring with early osteoarthritis have been suggested to decrease matrix osmolarity in situ (more hypotonic; see Ref. 35). During cartilage compression from daily activities, compaction of the cartilage matrix results in temporal changes in osmolarity (34). These changes are believed to contribute to changes in chondrocyte cell shape and volume in situ, potentially important parameters in mediating mechanotransduction (20, 63).

To gain a better understanding of the role that physical forces play in cartilage maintenance and degeneration, investigators have turned to cartilage explant studies that have yielded significant insights to the nature by which chondrocytes respond to applied mechanical loading. Deformational loading of cartilage explants has revealed distinct responses to regimes of static and dynamic loading. Because a plethora of physical stimuli exist in situ during tissue loading, contributions of concomitant stimuli, such as hydrostatic pressure, electrokinetic phenomena, and transport phenomena, have also been well investigated and have provided fundamental insights to how chondrocytes perceive their physical environment. Lesser understood, however, is the potential contributing factor of changes to the extracellular osmolarity that accompany tissue deformation, resulting from the alterations of the fixed charge density of the proteoglycan-rich extracellular matrix (18). A significant challenge toward achieving a better understanding of this phenomenon has been the inability for researchers to apply dynamic osmotic loads in culture. While static loading conditions facilitate experimental investigation, the natural physiological environment is dynamic and changes with time. In fact, studies have demonstrated that many cell types respond differently depending on whether a given mechanical stimulus (e.g., stretch, flow) is applied in a static or dynamic manner (1, 4, 12, 14, 36, 40, 44, 46, 58). For instance, while static deformation causes a reduction in chondrocyte biosynthetic activities, dynamic deformational loading results in an upregulation of chondrocyte biosynthetic activity (6, 49). Interestingly, while cyclic compression is innate to articular cartilage from daily activities, few studies to our knowledge have explored the response of cells to dynamic osmolarity changes.

Static hypotonic loading has been shown to change chondrocyte volume and intracellular pH, activate ion channels, and modulate cytoskeleton organization and aggrecan gene expression (13, 21, 27, 50, 51, 62). These studies have adopted manual solution changes or employed specialized chambers to achieve osmotic loading (24, 64). Although sufficient for static or even low-frequency experiments, these methodologies are not practical for the study of higher-frequency osmotic loading, and may be complicated by the shear effects that accompany fluid application. We have recently developed a novel microfluidic system to apply hydrostatic pressure-driven dynamic osmotic loading via composition modulated flow, where sequential exchanges of the same or different solutions can be applied to cultured cells (9). Moreover, this microfabricated flow device can achieve dynamic fluid exchanges with minimal fluid-shear stress (i.e., an order of magnitude smaller than generally observed to elicit a cellular response), and permit real-time monitoring of cellular responses using optical techniques. For the current channel geometry, the microfluidic device can apply osmotic loading of any magnitude up to a frequency of 0.1 Hz. The loading frequency is controlled by the interval between injection and withdrawal cycles, hence maintaining flow rate independent of loading frequency. This system may be applied in any kind of solution exchanges, such as growth factors and neurotransmitters. Using this microfluidic device, we have demonstrated the frequency dependent response of chondrocyte cell volume change to dynamic osmotic loading (9).

For the present study, we hypothesized that static and dynamic osmotic loading would result in differences in chondrocyte signal transduction, cytoskeleton organization, and gene expression. Cultured chondrocytes in the microfluidic device were subjected to dynamic hypotonic loading (310–110 mosM) from 0–0.1 Hz. The osmotic loading magnitude was chosen from the existing literature to accentuate the chondrocyte osmotic loading response, thereby facilitating study of and comparison of the chondrocyte response (cell size, intracellular calcium, cytoskeletal organization, and aggrecan gene expression) to applied dynamic osmotic loading with static osmotic loading (e.g., 5, 13, 56). The volume change and calcium signaling responses were recorded cell-by-cell using real-time fluorescence microscopy. Chondrocyte cytoskeletal organization and aggrecan and type II collagen gene expression were examined after 1 and 2 h of dynamic hypotonic loading, respectively. Chondrocytes exhibited increased calcium signaling with decreased loading frequency, with a positive correlation between cell volume and percentage of cells with calcium responses. At 0.1 Hz (sequential exchanges of 310 and 110 mosM solutions every 5 s), chondrocytes exhibited an actin cytoskeleton distribution more similar to isotonic control than that after static hypotonic loading. Aggrecan gene expression was stimulated in chondrocytes subjected to 0.1 Hz hypotonic loading.


   METHODS AND MATERIALS
TOP
 ABSTRACT
 METHODS AND MATERIALS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Microfluidic device. To achieve dynamic osmotic loading, a previously described microfluidic device was modified for biological studies (9, 54). In short, a Y-shaped microchannel system (Fig. 1A) was fabricated by replica-molding polydimethylsiloxane (Sylgard 184 kit, Dow Corning) from master structures. Before use, the microfluidic chip (polydimethylsiloxane Y-channel) was sterilized with ethanol and positioned onto a sterile glass coverslip (Fisher Scientific), which served as a base to seal the chip, forming closed rectangular channels (350 µm wide and 100 µm high) with open reservoirs at each end. The length of each Y-branch was ~1 cm. Composition modulated flow in the downstream branch was achieved by alternating fluid flow from each upstream reservoir (Fig. 1A, wide arrows). A hydrostatic pressure driven flow was obtained by two interlinked syringe pumps with an alternating fluid packets injection scheme. To confirm the composition flow modulation, FITC solutions (Sigma) were used to monitor flow patterns in the channel (Fig. 2).


Figure 1
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  		Fig. 1. A: microfluidic device, where {downarrow} indicates upstream reservoirs and <- indicates flow direction. B: sample chondrocyte volume normalized to initial cell volume (at time 0) and intracellular calcium response to dynamic hypotonic loading at 0.0167 Hz (9). F340/F380, fluorescence ratio at 340 and 380 nm excitation. Labels next to the hypotonic cell volume indicate peak volumes, and average peak-to-peak cell volume changes are calculated as

Formula 1

 

Figure 2
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  		Fig. 2. A: time-lapsed (1 and 0.1 Hz) fluorescence image of solution changes at 0.025 and 0.25 Hz in the microfluidic device. The y-axis represents the entire width of the channel and x-axis is time. B: normalized fluorescence intensity of the top, center, and bottom regions of the channel width image.

 
Cell culture. Articular cartilage was harvested from the wrist joints of freshly killed 4- to 6-mo-old calves. Primary chondrocytes were isolated via enzymatic digestion and plated at high cell density (~100,000 cells/cm2) for 2–4 wk, as previously described (37). Unless otherwise noted, all cell culture medium and reagents were obtained from Mediatech. For all studies, Dulbecco's modified essential medium supplemented with 10% fetal bovine serum (Sigma), amino acids (0.5x minimal essential amino acids, 1x nonessential amino acids), buffering agents [10 mM HEPES, 10 mM sodium bicarbonate, 10 mM TES, and 10 mM N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid], and antibiotics (100 U/ml penicillin, 100 mg/ml streptomycin) was used. The osmolarity of the media were determined to be 310 mosM with the use of a freezing-point osmometer (3D3 osmometer, Advanced Instruments).

General osmotic loading protocol and real-time calcium imaging. Before experiments, cells were released from the tissue culture dish using a 0.025% trypsin/EDTA solution, introduced to the Y-microfluidic channel and allowed to initiate adhesion onto 2 µg/ml poly-L-lysine-coated base (Sigma) of the channel for 45 min. Cells were then loaded with the calcium indicator dye fura-2 AM (5 nM; Molecular Probes). After incubation for 45 min, cells were equilibrated in serum-free isotonic Hanks' balanced salt solution (HBSS; Sigma). The microfluidic device was then placed on a precisely leveled inverted microscope stage (model IX-70, Olympus) at room temperature (~25°C). The two input and one output reservoirs were emptied. In each input well, a 50-µl aliquot of each solution was added and connected to a syringe pump (New Era Pump Systems) via a blood collection needle (21G; Becton-Dickinson). Experimental solutions were made by adding deionized water to HBSS to prepare hypotonic solution and the resulting osmolarity was confirmed with the osmometer. Dynamic hypotonic loading alternating between 310 and 110 mosM (or 310 and 310 mosM for isotonic control) was applied at frequencies of 0 (step loading), 0.0125, 0.0167, 0.025, 0.05, and 0.1 Hz. Fluorescence images were acquired using a x20 objective and charge-coupled device camera (Photon Technology International) controlled by MetaFluor software (Universal Imaging) using 340, 360, and 380 nm excitation wavelengths at >0.125 Hz for 10 loading cycles (0.335 µm/pixel image resolution). With the use of MetaFluor, individual cell regions were traced and their respective intracellular calcium levels expressed as a ratio of the 510 nm emission fluorescence intensities at 340 nm (calcium bound dye) and 380 nm (unbound dye), F340/F380 (16). A baseline calcium level for each cell was established by averaging the calcium ratio over the first 30 s before the start of static or dynamic osmotic loading described above. With the use of a "moving window," a custom-designed Matlab (The MathWorks) program was developed to identify "peaks" in the calcium levels. A calcium peak (or transient) was defined as a calcium increase of >3 x the standard deviation above the baseline level. Some cells exhibited multiple such peaks and were defined as repeat responders. Images acquired at the calcium-insensitive isosbestic point (360 nm) were used to monitor cell size changes (3).

Image processing and cell volume quantification. Cells in each field of view were tracked and the size of individual cells, measured in pixel area, was determined for each time point using a custom-designed Matlab program. The automated image processing algorithm included: 1) smoothing with a Gaussian filter, 2) computation of a bimodal intensity histogram, 3) identification of the "best" gray level threshold to isolate fluorescently labeled cells from background, and 4) binary segmentation via thresholding. Images from each time point were segmented individually and each cell detected in the image was assigned a label. To characterize cell size change in response to osmotic loading, cell volume was calculated from the pixel area (assuming a spherical geometry) and normalized to its respective initial isotonic size. Accuracy of the segmentation program and the spherical assumption were confirmed and reported in (9). Peak-to-peak cell volume changes were calculated by identifying the average changes between the maximum and minimum volume using a custom Matlab program (see Fig. 1 and Ref. 9).

Cytoskeletal organization. Effects of static and dynamic osmotic loading on chondrocyte cytoskeletal organization were assessed by labeling the actin cytoskeleton. Chondrocytes were subjected to static or dynamic hypotonic loading (310 to 110 mosM, 0.1 Hz) for 1 h and fixed with 3.7% formaldehyde. The actin cytoskeleton was labeled with 0.1 µM phalloidin (Oregon Green 488, Molecular Probes). Fluorescence images were acquired with an Olympus FluoView confocal microscope using a x60 oil-immersion objective. Quantitative analysis of F-actin distribution was conducted by measuring the intensity profile (average of two perpendicular diameters) through the center of the cell by using NIH Image J software and by normalizing the intensity of the cortical and plasma regions to the nuclear region (see Fig. 3) (13).


Figure 3
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  		Fig. 3. Scheme of quantification for cytoskeleton organization. Top, phalloidin staining for a sample chondrocyte F-actin is analyzed by two lines (5 pixels in width) drawn across the cell diameter. The average fluorescence profile is measured using Image J software and the cell edge is determined manually. The cell diameter was divided into six sections; the outer two cortical (C), middle two plasma (P), and the center two nuclear (N) regions. Fluorescence intensities of the three regions were averaged and normalized to the nuclear region.

 
Gene expression. To study the effects of static and dynamic osmotic loading on chondrocyte gene expression, cultured chondrocytes were plated at 2 million cells/ml in the microfluidic system for 45 min and subjected to osmotic loading for 2 h. Cells were harvested with TRIzol reagent (Invitrogen) and total RNA from each experiment was reverse transcribed with SuperScript III reverse transcriptase (Invitrogen). GAPDH, type II collagen, and aggrecan levels were quantified using an iCycler real-time PCR system and SYBR green supermix (Bio-Rad). Primers were designed with Beacon Designer (Premier Biosoft) and Primer3 online software (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi; see Ref. 48) using cDNA sequences obtained from National Center for Biotechnology Information Entrez Nucleotide web site (Table 1). Duplicates of 20-µl reactions were performed for each sample. Aggrecan and type II collagen mRNA levels were calculated using the –{Delta}{Delta}CT method (33) that normalized to the GAPDH and isotonic baseline levels.


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  		Table 1. Primers for real-time PCR

 
Pharmacological studies. Pharmacological inhibitors of calcium mobilization were studied for the possible different mechanisms involved in calcium signaling from static and dynamic hypotonic loading. Chondrocytes were cultured as described above and subjected to static and dynamic (0.025 and 0.1 Hz) hypotonic loading at 110 mosM. Calcium-free solutions were made with calcium-free HBSS and 10 mM EGTA (Sigma). Thapsigargin (1 µM; TG)-treated chondrocytes were incubated for 30 min before the experiments. TG is an inhibitor of intracellular calcium release from the endoplasmic reticulum.

Statistical analysis. All experiments were repeated at least twice and data were analyzed using Student's t-test or ANOVA with Fisher's post hoc test. {chi}2-tests were used to analyze the percentage of responding cells in the calcium response experiments. Pearson's test was used to test the correlation between cell volume and calcium responses. All analyses were done in Statistica (Statsoft) with {alpha} = 0.05. All error bars represent one standard deviation from the mean.


   RESULTS
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 ABSTRACT
 METHODS AND MATERIALS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Composition-modulated flow in the microfluidic channel is dependent on parameters such as channel width (W), average flow velocity (U), and applied modulation frequency (f). From these parameters, a dimensionless frequency (f*) can be found (see Ref. 55)

Formula 1(1)
Tang and co-workers have demonstrated that complete solution exchange (across the channel width) is achieved when the dimensionless frequency is <0.1, assuming a constant low Peclet number (54, 55). In the current channel geometry (W = 350 µm) and flow velocity (U = 750 µm/s), complete solution modulation can be achieved when applied modulation frequency is <0.21 Hz. In the range of 0.21–2.1 Hz (corresponding to 0.1 < f* < 1), fluid modulation occurs in the central region of the channel where the edges see only one solution type (54, 55). As shown in Fig. 2A, when one of the solutions is labeled with a fluorescent dye, dynamic modulation at 0.025 Hz resulted in complete fluid exchange across the channel width, whereas dynamic modulation at 0.25 Hz occurs in the center region of the microfluidic channel. Quantification of the fluorescence intensity confirms this observation (Fig. 2B).

Chondrocytes exhibited calcium transients in response to static (0 Hz) and dynamic hypotonic loading <0.05 Hz. Calcium transients and cell volume change of a representative cell are illustrated in Fig. 1B, which showed four calcium peaks and a peak-to-peak cell volume change of 0.29 (9). Chondrocytes also exhibited frequency-dependent cell size changes (Fig. 4A; n = 79–251) (9), whereas static loading resulted in an equilibrium volume change of 0.48 ± 0.26 (n = 73, 0 Hz). In addition, chondrocytes displayed frequency dependent calcium signaling (Fig. 4B). For cells exhibiting multiple calcium transients, these peaks appeared to coincide with the hypotonic loading phase of the dynamic osmotic loading (e.g., Fig. 1B). Interestingly, between the loading frequencies 0.0125–0.025 Hz, significantly more cells responded with multiple calcium peaks (n = 203–489; P < 0.05). No significant differences were observed for cell volume or calcium signaling in isotonic loading controls. A positive correlation was found between equilibrium cell volume change (for static hypotonic loading) and percentage of cells responding with calcium transients (Fig. 5A; n = 74, R = 0.651, P = 0.087). The magnitude of the equilibrium cell volume also exhibited a positive correlation with the magnitude of calcium peaks (Fig. 5B; n = 24, R = 0.695, P = 0.00016).


Figure 4
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  		Fig. 4. Effect of dynamic hypotonic loading frequency on cell volume (A) and calcium signaling (B). A: peak-to-peak volume changes were calculated by identifying the average changes between the maximum and minimum volume (n = 18–251 from >6 experiments, *P < 0.001 with all other frequencies and the isotonic control). B: percentage of chondrocytes that responded with calcium transients to dynamic hypotonic loading (n = 203–489 from >6 experiments). *P < 0.05 for all calcium responders vs. all others; §P < 0.004 for repeated calcium responders with that at frequencies of 0.1, 0.05 and 0 Hz; {dagger}P < 0.02 for all calcium responders with frequencies 0.025–0 Hz.

 

Figure 5
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  		Fig. 5. A: scatter plot of a histogram between the percentage of chondrocytes with calcium responses and equilibrium cell volume change for chondrocytes subjected to static hypotonic loading (n = 74 from 4 experiments, R = 0.637, P = 0.087). B: scatter plot of calcium peak magnitude change [(peak magnitude – baseline)/baseline] and equilibrium cell volume (n = 24, R = 0.695, P = 0.00016).

 
Examination of the actin cytoskeleton revealed that chondrocytes subjected to dynamic hypotonic loading for 1 h exhibited different cytoskeletal organization compared with cells subjected to isotonic or static hypotonic loadings. Confocal microscopy images of phalloidin staining demonstrated that chondrocytes under isotonic control exhibited cortical distribution of the actin cytoskeleton, whereas static hypotonic loading showed a trend of nuclear actin cytoskeleton localization (Fig. 6A). Quantitative analysis of the F-actin staining fluorescent intensity distribution revealed a similar trend, where dynamic hypotonic loading resulted in a more uniformly distributed actin cytoskeleton (Fig. 6B; n = 19–27).


Figure 6
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  		Fig. 6. A: confocal images of F-actin distribution in chondrocytes subjected to isotonic, static and dynamic (0.1 Hz) hypotonic loading for 1 h. Quantification of the actin fluorescence profile of 5 representative cells is plotted with the thick line representing the cell in picture above. Scale bar = 10 µm. B: quantitative analysis of F-actin distribution (see Fig. 3) (n = 19–27 from 4 experiments). *P < 0.05 with other corresponding cortical intensities.

 
The effect of dynamic osmotic loading on chondrocyte gene expression is shown in Fig. 7. Aggrecan and type II collagen expression are expressed as their mRNA levels normalized by GAPDH mRNA levels and the respective message levels of the isotonic controls. Dynamic hypotonic loading at 0.1 Hz for 2 h stimulated aggrecan gene expression by 49%, whereas static hypotonic loading suppressed aggrecan gene expression by 37% (Fig. 7A; n = 6–10, P = 0.018). Dynamic hypotonic loading at 0.1 Hz also resulted in increased type II collagen gene expression (Fig. 7B, n = 8–10).


Figure 7
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  		Fig. 7. Chondrocyte relative gene expression of aggrecan (AGC; A) and type II collagen (COLII; B) in response to static and dynamic hypotonic loading at 0.1 Hz (*P < 0.02 compared with 0.1 Hz; n = 6–10 from 2 experiments). Real-time PCR analysis using primers listed in Table 1 and aggrecan and type II collagen mRNA levels were normalized to that of GAPDH and isotonic controls.

 
To study the mechanisms involved in calcium signaling from static and dynamic osmotic loading, chondrocytes were subjected to calcium-free EGTA hypotonic loading to eliminate extracellular calcium source or treated with TG, an intracellular calcium release inhibitor. Both EGTA and TG significantly suppressed the percentage of cells responding with a calcium transient to static hypotonic loading, with EGTA exerting a larger diminution in the percentage of cells responding (Fig. 8; P < 0.05). In dynamic hypotonic loading, no changes in responding cells were observed for either EGTA or TG (n = 135–213 cells).


Figure 8
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  		Fig. 8. Differential calcium signaling mechanisms in static (0 Hz) and dynamic hypotonic loading. *P < 0.05 in overall percentage of calcium responders compared with the static groups at the same loading frequency, {dagger}P < 0.01 in percentage of repeat responders (cells with multiple calcium transients) with the static groups at the same loading frequency, n = 135–213 cells.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS AND MATERIALS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In this study we have demonstrated that cultured articular chondrocytes exhibited a frequency-dependent intracellular calcium transient response to applied dynamic osmotic loading. The experiments (dynamic hypotonic loading from 0 to 0.1 Hz) were performed using a novel microfluidic device with minimal concomitant shear stress. The latter is supported by a lack of calcium response observed in control chondrocytes subjected to dynamic exchanges of isotonic media. This calcium response is observed to correlate with magnitude of chondrocyte volume change (Fig. 5). Chondrocytes have been reported to behave as perfect osmometers where their equilibrium volume correlates with the applied osmotic load (17, 56). Chondrocyte cell size change was inversely related to applied osmotic loading frequency, as previously reported by our laboratory (9). This finding suggests that this cell size change reflects the duration of osmotic loading exposure (i.e., 1/frequency). Taken together with our findings in Fig. 5, the triggering of calcium transients in response to hypotonic loading may be related to the extent of volume/membrane expansion (17). Calcium is one candidate signaling molecule that has been shown to be important in chondrocyte mechanotransduction and can be monitored during stimulus application (2, 13, 38, 39, 47, 62, 64).

Examination of the actin cytoskeleton revealed differential organization for chondrocytes subjected to dynamic osmotic loading compared with cells subjected to isotonic or static hypotonic loadings (Fig. 6). Erickson and co-workers (13) previously reported that static hypotonic loading of chondrocytes results in calcium-induced actin reorganization. This reorganization may be required for the delayed volume regulatory decrease responses (time constant, ~8 min; see Refs. 7 and 45). Following up the hypotonic loading response with the response to the isotonic "recovery" phase in dynamic osmotic loading may contribute to the differences in the calcium response and cytoskeleton organization for dynamic osmotic loading compared with that for static osmotic loading. Because the actin cytoskeleton has been proposed as a mechanosensor in various cell types (22, 23, 61), this differential actin reorganization between static and dynamic osmotic loading may lead to downstream secondary changes in cell signaling and activities (such as gene expression and cell biosynthesis, e.g., Ref. 4).

Under the experimental conditions prescribed in our studies, dynamic osmotic loading was found to increase chondrocyte aggrecan gene expression (Fig. 7). The osmotic loading magnitude for the current study was chosen from that studied in the chondrocyte literature (7, 13, 27, 50, 62), in part to facilitate comparison of our novel dynamic osmotic experiments with those of static loading by other researchers. This finding demonstrates that dynamic osmotic loading has a similar stimulatory effect on chondrocytes as other applied dynamic mechanical stimuli. Dynamic deformational loading, for example, increases aggrecan and collagen production in cartilage explants and chondrocytes encapsulated in three-dimensional hydrogels (6, 28, 36, 49). A relationship between intracellular calcium and chondrocyte aggrecan gene expression has been reported by Alford and co-workers (2). General calcium inhibitor studies showed different calcium entry pathways in static and dynamic hypotonic loading (Fig. 8). While we may speculate that dynamic hypotonic loading may modulate chondrocyte aggrecan gene expression through differential intracellular calcium signals, further investigations are required to establish whether such a direct link exists.

We apply dynamic osmotic loading via cyclic alterations of the extracellular osmolarity, as verified with our fluorescence dextran labeling protocol (see Fig. 2 and Ref. 9). It is difficult to know whether the chondrocyte perceives an osmotic pressure (resulting from the alterations in extracellular osmolarity) because there are no techniques that permit the osmotic pressure inside a cell to be measured. However, the degree of water flow across the cell membrane (in response to a differential solute gradient) ultimately depends on its hydraulic permeability. Osmotic pressure can be empirically determined using Kedem-Katchalsky equations that equate cell volume change with osmotic pressure (at equilibrium) (30, 31). For static loading experiments, it is clear that the cells are seeing an osmotic gradient across the cell membrane with a corresponding development of osmotic pressure, as an obvious cell size change is observable and is consistent with the Kedem-Katchalsky model. We argue that the faster the exchange of solutions, the less time there is for water to traverse the cell membrane, and that accordingly less osmotic pressure develops. The osmotic pressure developed in the cell under dynamic osmotic loading represents a fraction (as noted by the maximum cell size that is reached) of the maximum osmotic pressure (that is directly equal to the applied osmotic loading) observed in static osmotic loading. Ultimately, our results are interesting in that the chondrocyte appears to be responsive to exposure to the osmotic gradient (despite lack of measurable cell size change) and further studies are required to delineate between the solute concentration gradient (between extracellular and intracellular levels) and intracellular osmotic pressure changes.

Advances in microfluidics have provided the opportunity for development of novel systems for biological applications, such as cell patterning and nanoscale assays (e.g., 10, 29, 53, 59, 60). The current application utilizes a novel microfluidic flow chamber developed by our laboratory that permits solutions of alternating composition (i.e., cyclic osmotic loading) to be applied to cultured cells, and with the capability of permitting cell measurements using optically based techniques and probes (e.g., green fluorescent protein constructs, ion-sensitive probes, and optical laser tweezers). With the current design, a complete dynamic fluid modulation (where the solution across the channel width is completely exchanged) at 0.1 Hz was studied. By confining the field of view for analysis (to the central region) or optimizing the channel geometry, dynamic solution exchanges can be achieved at a higher rate (Fig. 2 and Eq. 1) (25). Alternatively, a range of dynamic osmotic loading magnitudes (dependent on location relative to the channel center or wall) at higher frequencies can also be studied in the same experiment by analyzing the cell response at different locations (with respect to the channel width direction).

We have previously demonstrated the feasibility of the microfluidic system for dynamic osmotic loading applications and reported the frequency dependence of chondrocyte volume changes (9). In the current investigation, we extended these studies to explore biological responses of chondrocytes to dynamic hypotonic loading-induced calcium signaling, cytoskeleton organization and gene expression. Dynamic hypotonic loading results in frequency-dependent calcium signaling, altered cortical actin organization, and stimulated chondrocyte aggrecan gene expression. Future studies will aim to determine the direct relationship between the observed intracellular calcium response and cytoskeletal re-organization on aggrecan gene expression. Support for the aims of our study is provided by a recent investigation by Szafranski and co-workers (52) who found chondrocyte organelle volume changes from mechanical deformation of cartilage and proposed that these changes arise from osmotic forces. They suggest that the deformation of chondrocyte organelles contribute to mechanotransduction pathways linking translational and posttranslational events to cell deformation. Elucidation of mechanisms that mediate chondrocyte mechanotransduction to dynamic stimuli may help to explain the role of physical factors in the etiology and progression of osteoarthritis, and contribute to efforts that apply physiologic loading for in vitro development of engineered tissue for cartilage repair (8).


   GRANTS
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 ABSTRACT
 METHODS AND MATERIALS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was funded by National Institutes of Health Grants AR-48791 and AR-052871 and by the Whitaker Foundation.


   ACKNOWLEDGMENTS
 
We thank Lawrence David, Jake Hoffman, and Dr. Chris Wiggins for the use of the image thresholding program.


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. T. Hung, Dept. of Biomedical Engineering, 351 Engineering Terrace, 1210 Amsterdam Ave., MC 8904, New York, NY 10027 (e-mail: cth6{at}columbia.edu)

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.


   REFERENCES
TOP
 ABSTRACT
 METHODS AND MATERIALS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
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