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(Radiology. 2001;220:428-435.)
© RSNA, 2001

Experimental Studies

In Vitro Effects of Transcatheter Injection on Structure, Cell Viability, and Cell Metabolism in Fibroblast-impregnated Alginate Microspheres1

Todd Abruzzo, MD, Harry J. Cloft, MD, PhD, George G. Shengelaia, MD, Sandra Miller Waldrop, PhD, David F. Kallmes, MD, Jacques E. Dion, MD, Ioannis Constantinidis, PhD and Athanassios Sambanis, PhD

1 From the Section of Interventional Neuroradiology (T.A., H.J.C., G.G.S., J.E.D.) and Division of Radiological Sciences (S.M.W., I.C.), Department of Radiology, Emory University School of Medicine, 1364 Clifton Rd NE, Atlanta, GA 30302; School of Chemical Engineering and P. H. Parker Institute for Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta (A.S.); and Division of Interventional Neuroradiology, Department of Radiology, University of Virginia Health Sciences Center, Charlottesville (D.F.K.). Received August 3, 2000; revision requested September 19; revision received December 19; accepted January 26, 2001. Supported by a grant from the Emory University/Georgia Institute of Technology Biomedical Research Center. H.J.C. supported in part the RSNA Research and Education Foundation as a 1999 Sterling Diagnostic/RSNA Scholar. Address correspondence to T.A. (e-mail: tabruzz@emory.edu).


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
 
PURPOSE: To determine if microsphere-encapsulated cell preparations can be delivered through a microcatheter without compromising microsphere structure, cell viability, or metabolism.

MATERIALS AND METHODS: Fibroblast-impregnated microspheres were fabricated by using 1.0% alginate and rabbit synovial fibroblasts. Fibroblast-impregnated alginate microspheres injected through microcatheters were analyzed in parallel with identical noninjected microspheres. The effects of transcatheter injection on structure and cell viability (percentage of viable cells per microsphere) were correlated with microsphere size. Structural effects were analyzed by using light microscopy, and 7-day percentage (ratio of live cells to dead cells) cell viability was assessed with confocal microscopy and fluorescent staining. In a second series of experiments, the metabolism of small microspheres was studied during a course of 7 days by using a spectrophotometric bioanalyzer.

RESULTS: Transcatheter injection caused fracturing and/or fragmentation of large (800–1,000 µm) and medium (500–750 µm) microspheres, while small (250–400 µm) microspheres were structurally unaffected by transcatheter injection. Fracturing and fragmentation were associated with cell release from the alginate matrix. Although transcatheter injection reduced cell viability by 17%–23% in all size categories, it did not cause a detectable alteration in the rate of glucose metabolism.

CONCLUSION: Transcatheter injection was physiologically well tolerated by fibroblasts encapsulated in alginate microspheres; however, when microsphere diameter exceeded the catheter diameter, fracturing and fragmentation of microspheres compromised the sequestration function of the microsphere vector.

Index terms: Microspheres • Experimental study


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
 
The implantation of therapeutically active cells into diseased organs and tissues represents a promising treatment modality. Alginate hydrogels have evolved as a form of cell entrapment medium for the immunologic and physical isolation of cell grafts (1–4). Alginates are polysaccharide chains composed of linked mannuronic and guluronic acid residues (2). In solution, individual chains can be made to form a hydrogel through the interchain chelation of calcium or other divalent cations (2). While cells entrapped in alginate remain physically isolated from the surrounding tissues, they are able to influence tissue structure and function through the release of diffusible mediators. Surgically implantable alginate microspheres bearing genetically engineered cells have been used to accomplish sustained delivery of insulin, growth hormone, and factor IX in experimental animals (2,5–12). Such microspheres are similar in size to currently used particle embolic agents such as polyvinyl alcohol (13).

Transcatheter injection may be an alternative nonsurgical approach to the delivery of such constructs. Embolization of a small or medium-sized artery with alginate-encapsulated cell preparations may enable the establishment of endovascular cell allografts (14). In the absence of a vigorous foreign body reaction, endovascular cell allografts may recruit a blood supply from the vaso vasorum of the host vessel and persist in a functional state.

Since injection through a microcatheter exposes embolic agents to severe mechanical forces, it is conceivable that transcatheter injection might have an adverse effect on microsphere structure, cell viability, and cell function, which precludes it as a potential delivery scheme. The following in vitro investigation was performed (a) to establish the feasibility of delivering alginate entrapped cells with a microcatheter and (b) to evaluate the effects on cell metabolism of microcatheter injection of alginate microspheres impregnated with living cells.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
 
Fabrication of Alginate Microspheres
Sterile filtered (0.2-µm pore size) solutions of 1.0% low-viscosity high-mannuronic alginate (Pronova, Lysaker, Norway) were prepared by dissolving it in 0.9% sodium chloride. The lowest technically feasible alginate concentration was used because we believed that maximizing hydrogel fluidity would minimize the resistance encountered by microspheres during catheter transit. Findings from our preliminary investigations suggested that structurally stable microspheres could not be consistently fabricated from lower alginate concentrations (<=0.5%).

Microspheres containing approximately 106 cells per milliliter were prepared by using rabbit synovial fibroblasts (HIG-82; American Tissue Culture Corporation, Manassas, Va) cultivated in medium (Dulbecco modified essential medium; Mediatech, Herndon, Va) supplemented with 10% fetal bovine serum. Confluent fibroblast monolayers were treated with trypsin, pelleted with centrifugation, and washed twice with serum-supplemented medium. Cell pellets were mixed in 1% filtered low-viscosity high-mannuronic alginate to obtain alginate cell suspensions.

Alginate microspheres were prepared by hand injecting alginate cell suspensions through a pressure-regulated airflow microbead generator into a Petri dish containing sterile filtered 1.1% calcium chloride (3,15). Small microspheres were created by using a generator pressure of 15 psi and passing the resulting product through a sieve (Phosphor Bronze; Dual MFG, Chicago, Ill) with 200 openings per linear inch (127-µm pore size). Medium and large microspheres were created with generator pressures of 12.5 and 10.0 psi, respectively. The calcium chloride solution was aspirated, and the microspheres were washed several times with serum-supplemented medium.

Microspheres were incubated in serum-supplemented medium for 24 hours, then washed with serum-free medium. For each population of microspheres (small, medium, large), size was confirmed by two of the authors (T.A., G.G.S.) using sample diameter measurements on a hemacytometer slide with a light microscope. The reported range of measurements for each size category was obtained by scanning the sample. The smallest and largest microspheres in the sample were measured until the upper and lower limits of an individual size category were defined.

All experiments were performed with freshly prepared fibroblast-impregnated microspheres in serum-free medium to achieve a relatively stable nonproliferating population of cells in each microsphere.

Analysis of Microsphere Structure and Cell Viability
Three populations (small, medium, large) of fibroblast-impregnated alginate microspheres were prepared and divided into control and transcatheter groups on day 1. For each population, a 0.75-mL aliquot of control microspheres was transferred to one well of a multiwell plate, and 3 mL of serum-free medium was added. The approximate number of control microspheres was 4.17 x 104 in the small-size group, 5.87 x 103 in the medium-size group, and 1.96 x 103 in the large-size group.

For each population, an additional 0.75 mL of microspheres, suspended in 5 mL of serum-free medium, was loaded into a 5 mL syringe. The syringe contents were manually injected through a microcatheter (Tracker 18; Boston Scientific, Natick, Mass), 150 cm in length, into a second well of the multiwell plate. The contents of each catheter were flushed into the corresponding well with an additional 5 mL of serum-free medium, and excess medium was aspirated until a residual volume of 3 mL was present. A single injection was performed for each size category (a total of three injections), and the multiwell plate was then placed on a rotary shaker in an incubator at 37°C. All three injections were performed by the same author (T.A.), and catheter resistance to microsphere injection and injection difficulty were subjectively assessed. The approximate number of injected microspheres was 4.17 x 104 in the small-size group, 5.87 x 103 in the medium-size group, and 1.96 x 103 in the large-size group.

Starting on day 1, microsphere structure was analyzed (T.A., G.G.S.) by means of daily examination of the multiwell plates with a light microscope. Microsphere structure was analyzed to determine the presence of structural flaws (erosions, fragmentation, or fracturing). Cell release was assessed by inspecting the dependent surfaces of the wells for free-growing fibroblasts. Structural stability was assessed by noting changes in structural integrity throughout the 7-day course of the experiment. On day 7, all microspheres were collected and distributed into six conical tubes, each representing a size category (small, medium, and large) of transcatheter or control microspheres. The contents of each tube were then treated with a fluorescent stain (Molecular Probes, Eugene, Ore), which consisted of calcein acetoxymethyl ester and ethidium homodimer-1, to differentiate between the live and dead cells.

The stained microspheres were then analyzed by using confocal microscopy with an argon laser (Carl Zeiss, Thornwood, NY). Cells fluorescing red were counted as dead, since red fluorescent ethidium can penetrate and intercalate into the nucleic acids of only cells with damaged cytoplasmic membranes (16). Cells fluorescing green were counted as viable, since only viable cells can enzymatically convert the nonfluorescent membrane permeant calcein acetoxymethyl ester to nonpermeant green fluorescent calcein (16). Eight representative microspheres from each tube were randomly selected for cell viability determinations.

Random selection of microspheres was accomplished by vortex mixing each tube just prior to micropipetting a 50-µL sample from the tube onto a glass slide. Two microspheres were randomly selected from each quadrant of the glass slide for analysis. The total number of microspheres distributed in each quadrant of a slide was approximately 10 for large microspheres, 37 for medium microspheres, and 260 for small microspheres. Each sample was considered representative of its parent population due to the random distribution produced by vortex mixing.

Each selected microsphere was scanned by using a standard confocal technique to generate a stacked series of cross-sectional images, each representing a 6-µm-thick section of the microsphere.

Two median and two paramedian image sections were selected for further analysis. Cell viability was calculated as the ratio of green fluorescence (live cell) surface area to the combined surface area of green fluorescence (live cells) and red fluorescence (dead cells). Representation in a surface-area summation was based on a fluorescence-intensity threshold that was set to the most weakly fluorescent cell in the image being analyzed. Dead cell fragments and debris were, therefore, proportionately represented in the analysis.

By using an image browser program (LSM 510; Carl Zeiss, Thornwood, NY), one of the authors (T.A.) drew a region of interest around the microsphere borders on the surface projection of each of the four representative slabs. By applying a series of masks to the image, the individual surface areas constituted by live cells (green fluorescence) and dead cells (red fluorescence) in the region of interest were determined (Fig 1). The ratio of live cells to total cells in each representative section was then approximated by a calculation: green fluorescence surface area/(green fluorescence surface area plus red fluorescence surface area). The average percentage of microsphere cells viable was calculated for each group by averaging the live-to-total cell ratio for four slabs in eight microspheres (n = 32).



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  		Figure 1a. Cell viability in microspheres. (Original magnification, x40.) (a) Photomicrograph of a transmedian confocal microscopic section through a 7-day medium transcatheter microsphere with a region of interest (white circle) around the microsphere perimeter. Since each section is 6-µm thick, volume averaging is eliminated (cell size, >=6 µm). Live cells (long arrows) fluoresce green, and dead cells (short arrows) fluoresce red. (b) Same image as in a with red fluorescence subtracted allows calculation of live cell surface area. (c) Same image as in a with green fluorescence subtracted allows calculation of the dead cell surface area. Note that in some areas, dead cells are fractured into multiple closely related fragments or minute particles of scattered debris. The fluorescence of subcellular fragments and dead cell debris (arrows) was proportionately represented in the total surface area summation for dead cells.

 


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  		Figure 1b. Cell viability in microspheres. (Original magnification, x40.) (a) Photomicrograph of a transmedian confocal microscopic section through a 7-day medium transcatheter microsphere with a region of interest (white circle) around the microsphere perimeter. Since each section is 6-µm thick, volume averaging is eliminated (cell size, >=6 µm). Live cells (long arrows) fluoresce green, and dead cells (short arrows) fluoresce red. (b) Same image as in a with red fluorescence subtracted allows calculation of live cell surface area. (c) Same image as in a with green fluorescence subtracted allows calculation of the dead cell surface area. Note that in some areas, dead cells are fractured into multiple closely related fragments or minute particles of scattered debris. The fluorescence of subcellular fragments and dead cell debris (arrows) was proportionately represented in the total surface area summation for dead cells.

 


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  		Figure 1c. Cell viability in microspheres. (Original magnification, x40.) (a) Photomicrograph of a transmedian confocal microscopic section through a 7-day medium transcatheter microsphere with a region of interest (white circle) around the microsphere perimeter. Since each section is 6-µm thick, volume averaging is eliminated (cell size, >=6 µm). Live cells (long arrows) fluoresce green, and dead cells (short arrows) fluoresce red. (b) Same image as in a with red fluorescence subtracted allows calculation of live cell surface area. (c) Same image as in a with green fluorescence subtracted allows calculation of the dead cell surface area. Note that in some areas, dead cells are fractured into multiple closely related fragments or minute particles of scattered debris. The fluorescence of subcellular fragments and dead cell debris (arrows) was proportionately represented in the total surface area summation for dead cells.

 
Metabolic Studies
A homogenous population of fibroblast-impregnated alginate microspheres, 250–400 µm in diameter, was used to determine the effect of transcatheter injection on cell metabolism. Only small microspheres were used for metabolic studies because larger microspheres were known to release cells after transcatheter injection. We believed that cell release would unpredictably bias the results of metabolic studies, since liberated cells freely growing in wells are known to grow and metabolize at rate faster than that of encapsulated cells (17,18).

Glucose consumption was studied because glucose is the primary energy substrate for oxidative metabolism in cultured eukaryotic cells, and changes in glucose metabolism mirror global changes in cellular function. Furthermore, accurate quantitative analysis of glucose consumption is easily performed. A 0.75-mL aliquot of control microspheres was transferred to one well of a multiwell plate, and 3 mL of serum-free medium was added. Another 0.75-mL microsphere aliquot was manually injected through a microcatheter (Tracker 18; Boston Scientific) into a second well by using the technique described in the previous section. Paired transcatheter and control specimen multiwell plates were prepared in triplicate (three independent trials performed on separate days by using separate alginate cell suspensions). Each plate was placed on a rotary shaker in an incubator at 37°C.

Initial (days 1–3) and delayed (days 3–7) rates of glucose consumption were measured by determining the glucose content in each well at selected time points (initial, 24 hours, 48 hours, 142 hours, 156 hours). On day 3, the residual medium in each well was exchanged for 3 mL of fresh serum-free medium (glucose content was determined before and after the medium exchange). In each case, the rate was calculated in milligrams per hour as the slope of a three-point scatterplot of media glucose content versus time. The glucose concentration of media samples (0.5 mL per sample) extracted from each well at the selected time points was determined by using a bioanalyzer (Ektachem DT60; Eastman Kodak, Rochester, NY).

Statistical Analysis
Cell viability in control and transcatheter microspheres was compared for each of the three size categories by using an analysis of variance and the Tukey procedure for multiple comparisons. A t test for paired data sets was used to analyze differences in glucose metabolism between control and transcatheter microspheres.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
 
Microsphere Structure
Microsphere diameter ranged from 800 to 1,000 µm in the large-size category (286–523 cells per microsphere), from 500 to 750 µm in the medium-size category (65–221 cells per microsphere), and from 250 to 400 µm in the small-size category (eight to 33 cells per microsphere) (Fig 2).



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  		Figure 2a. Photomicrograph of control microspheres of each size category shown on hemacytometer slides. (a) Large microsphere measuring approximately 1,000 µm in diameter. Distance between arrowheads is 250 µm. (Original magnification, x40.) (b) Medium microspheres measuring approximately 750 µm in diameter. (Original magnification, x40.) Distance between arrowheads is 250 µm. (c) Small microsphere measuring approximately 280 µm in diameter. Distance between arrowheads is 62.5 µm. (Original magnification, x100.)

 


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  		Figure 2b. Photomicrograph of control microspheres of each size category shown on hemacytometer slides. (a) Large microsphere measuring approximately 1,000 µm in diameter. Distance between arrowheads is 250 µm. (Original magnification, x40.) (b) Medium microspheres measuring approximately 750 µm in diameter. (Original magnification, x40.) Distance between arrowheads is 250 µm. (c) Small microsphere measuring approximately 280 µm in diameter. Distance between arrowheads is 62.5 µm. (Original magnification, x100.)

 


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  		Figure 2c. Photomicrograph of control microspheres of each size category shown on hemacytometer slides. (a) Large microsphere measuring approximately 1,000 µm in diameter. Distance between arrowheads is 250 µm. (Original magnification, x40.) (b) Medium microspheres measuring approximately 750 µm in diameter. (Original magnification, x40.) Distance between arrowheads is 250 µm. (c) Small microsphere measuring approximately 280 µm in diameter. Distance between arrowheads is 62.5 µm. (Original magnification, x100.)

 
There were no structural flaws detected in control microspheres of any size. The structure of control microspheres remained stable throughout the entire 7 days of the study, without evidence of fracture, fragmentation, erosion, or cell release.

Although no substantial resistance or difficulty was encountered during any of the transcatheter injections, large and medium transcatheter microspheres were substantially damaged immediately after injection. Large transcatheter microspheres were either fragmented into several amorphous fragments and granules or sustained multiple fractures (Fig 3). Fragmentation of microspheres resulted in the liberation of large numbers of fibroblasts from the alginate matrix immediately after transcatheter injection. These fibroblasts attached to the surface of their corresponding wells and appeared to grow in number throughout the 7-day course of the experiment. Medium transcatheter microspheres possessed one or two fractures each. Occasional fibroblasts were found growing on the surfaces of the wells containing medium microspheres; this finding indicated fibroblast release.



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  		Figure 3a. Photomicrograph of structural effects of transcatheter injection. (Original magnification, x40.) (a) Fractured large transcatheter microsphere. Arrows = fractures. (b) Fragmented large transcatheter microsphere. Straight arrows = fragmentation. Curved arrows = fibroblasts liberated from the microsphere.

 


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  		Figure 3b. Photomicrograph of structural effects of transcatheter injection. (Original magnification, x40.) (a) Fractured large transcatheter microsphere. Arrows = fractures. (b) Fragmented large transcatheter microsphere. Straight arrows = fragmentation. Curved arrows = fibroblasts liberated from the microsphere.

 
Small transcatheter microspheres demonstrated no evidence of structural compromise throughout the entire 7 days. Fibroblast release from small transcatheter microspheres was not detected.

Cell Viability
In comparison with control microspheres, cell viability was reduced in the transcatheter microspheres in all size categories (Fig 4). Transcatheter microspheres showed a statistically significant decrease in cell viability relative to that of control microspheres for all three categories (P < .001). The percentage reduction in cell viability was determined by using the following equation: [(average fraction of viable cells per control microsphere - average fraction of viable cells per transcatheter microsphere)/average fraction of viable cells per control microsphere] x 100.



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  		Figure 4a. Graph represents the effect of transcatheter injection on cell viability. The average fraction of total microsphere fibroblasts that were viable after day 7 is displayed. Transcatheter microspheres are represented by white columns and control microspheres are represented by black columns. Error bars indicate the average ± 2 SDs. Note that for each size category the average fraction of viable fibroblasts is slightly lower in the transcatheter group. (a) Small microspheres (250-400-µm diameter). (b) Medium microspheres (500-750-µm diameter). (c) Large microspheres (800-1,000-µm diameter).

 


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  		Figure 4b. Graph represents the effect of transcatheter injection on cell viability. The average fraction of total microsphere fibroblasts that were viable after day 7 is displayed. Transcatheter microspheres are represented by white columns and control microspheres are represented by black columns. Error bars indicate the average ± 2 SDs. Note that for each size category the average fraction of viable fibroblasts is slightly lower in the transcatheter group. (a) Small microspheres (250-400-µm diameter). (b) Medium microspheres (500-750-µm diameter). (c) Large microspheres (800-1,000-µm diameter).

 


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  		Figure 4c. Graph represents the effect of transcatheter injection on cell viability. The average fraction of total microsphere fibroblasts that were viable after day 7 is displayed. Transcatheter microspheres are represented by white columns and control microspheres are represented by black columns. Error bars indicate the average ± 2 SDs. Note that for each size category the average fraction of viable fibroblasts is slightly lower in the transcatheter group. (a) Small microspheres (250-400-µm diameter). (b) Medium microspheres (500-750-µm diameter). (c) Large microspheres (800-1,000-µm diameter).

 
The percentage reduction in cell viability was 23.3% ([0.2072/0.8902] x 100) in small microspheres, 17.3% ([0.1532/0.8843] x 100) in medium microspheres, and 21.7% ([0.1912/0.8807] x 100) in large microspheres. The magnitude of cell viability reduction did not correlate with microsphere size by using the Pearson correlation coefficient (P = .834). The Tukey test for multiple comparisons showed that the viability for a group of transcatheter microspheres (small, medium, or large) was significantly lower than the viability for control microspheres in any size group and not just the corresponding size group (P = .001). Additionally, the viability of a size of transcatheter or control microspheres was not significantly different from that of microspheres of other sizes in its group (transcatheter or control).

Cell Metabolism
The average initial rate of glucose metabolism (days 1–3) for the three trials was 0.15 mg/h for control microspheres and 0.12 mg/h for transcatheter microspheres. The average delayed rate of glucose metabolism (days 3–7) for the three trials was 0.091 mg/h for control microspheres and 0.090 mg/h for transcatheter microspheres. The average percentage difference in the initial rate of glucose metabolism between transcatheter and control microspheres for a trial was 17.591%, or [(0.008/0.079 + 0.030/0.170 + 0.050/0.200)/3] x 100 (Fig 5). The average percentage difference in the delayed rate of glucose metabolism between transcatheter and control microspheres for a trial was 3.385% [(0.000/0.073 + 0.010/0.110 + 0.001/0.094)/3] x 100 (Fig 5).



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  		Figure 5. Graph of the average difference in the rate of glucose metabolism between transcatheter and control microspheres as a percentage of the rate of glucose metabolism for control microspheres (control rate - transcatheter rate/control rate). The white column represents the initial (days 1-3) rate, and the black column represents the delayed (days 3-7) rate. Error bars indicate the average ± 1 SD of the three trials represented in each column. The relative depression of metabolic rate associated with transcatheter injection diminishes almost completely in several days.

 
By using the paired t test to compare transcatheter and control groups for each trial, there was no significant difference in the initial or delayed rate of glucose metabolism. A paired t test comparison of the initial and delayed rates for each trial showed no significant difference.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
 
Hydrogel Microspheres as Particulate Embolic Agents
Gobin et al (19) recently discussed the technical advantages of hydrogel microspheres as particulate embolic agents. Unlike other particulate embolics, such as polyvinyl alcohol, hydrogels can easily be made radiopaque by means of the incorporation of tantalum or tungsten powder. In a similar fashion, drugs can be incorporated into hydrogel embolics to confer therapeutic effects (20,21). Hydrogel microspheres possess further advantages because uniform particles of precise dimensions can be created, permitting predictable occlusion of specific small target vessels. Hydrogels have a low friction coefficient and do not clump together or adhere to surrounding tissues; these characteristics ensure that vessel occlusion will occur only when the particles become impacted in a vessel of the same size. In addition, hydrogel microspheres have a smooth surface and are easily deformable, allowing relatively easy injection through microcatheters. Alginate hydrogels are also biologically advantageous, since they have excellent biocompatibility and provoke minimal inflammatory response (22). The alginate hydrogel microspheres used in our experiments can potentially be used to deliver living cells into human tissues for the treatment of a variety of diseases.

Effects of Transcatheter Injection on Microsphere Structure
Structural defects compromise the sequestration function of alginate microspheres harboring cell grafts. In this study, we found evidence of cell release from fractured and fragmented microspheres. Cells released from the hydrogel matrix are free to migrate into host tissues and/or interact with immune-effector cells and molecules.

Although Gobin et al (19) reported that polyacrylonitrile microspheres as large as 700 µm in diameter were easily injected through a 0.018-inch microcatheter, they did not examine the structural consequences of this manipulation. Similar to them, we experienced no mechanical difficulty during injection, even with microspheres up to 1,000 µm in diameter; however, microspheres 500 µm in diameter or larger had substantial structural damage. This upper limit of microsphere diameter roughly approximates the 533-µm inner diameter of the Tracker 18 microcatheters (technical information provided by Boston Scientific) used in our experiments (Fig 6).



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  		Figure 6. Photomicrograph shows the physical constraints imposed by microcatheter dimensions. A 500-µm fibroblast-impregnated 1%-alginate microsphere is shown exiting the distal ostium of a microcatheter. Arrowheads outlining the tip of the catheter demarcate a distance of 533 µm.

 
Our results indicate that alginate microspheres harboring cell grafts should not be delivered through a microcatheter unless the cross-sectional diameter of the microcatheter is at least as big as that of microsphere. Although increasing the concentration of hydrogel matrices may increase resistance to fracturing, it may also adversely compromise cell growth and function (23–25). Alginate preparations with a high guluronic acid content, in contrast to the preparation with a high mannuronic content used in this study, may also increase microsphere strength. However, prior study findings have demonstrated that such alginates have a negative effect on cell growth and the overall metabolic activities of microspheres (26).

In general, hydrogel modifications that augment mechanical strength by increasing rigidity and decreasing fluidity inhibit cell growth and may actually cause excessive cell death in the matrix (2,26). Microsphere coating with poly-L-lysine has been shown to increase microsphere stability without compromising cell function, and this may be a useful modification to prevent cell release from hydrogel microspheres intended as cell-carrier embolic agents (17). Although it stands to reason that larger catheters (>=5 F) would permit the injection of larger microspheres without causing fracturing or fragmentation; additional research is necessary to establish the effect of catheter diameter and length on microsphere structure.

Effect of Transcatheter Injection on Cell Viability and Metabolism
In this study, transcatheter injection caused a small but statistically significant decrease in the percentage of viable cells per microsphere. The magnitude of this effect did not correlate with microsphere size. Several mechanisms of mechanical cell injury are plausible. In addition to acceleration and deceleration, cells are also subjected to compression forces during catheter transit. Although the positive pressure gradient experienced by fibroblasts passing through a microcatheter may increase with microsphere size, there may be a threshold effect on cell physiology so that the percentage of cell loss is the same for microspheres of all sizes. Mechanical forces are known to influence cell physiology by means of changes in cytoskeletal structure, membrane function, and gene expression (27,28). Some study findings (29,30) have demonstrated that mechanical forces, especially compression, can result in apoptotic cell death.

Although mechanical stresses may have contributed to cell death in this study, a more likely cause of the observed loss of cell viability is the chemical stress experienced by the cells during catheter transit. Clearly, the cells were not in an ideal culture environment during this procedure. For instance, hypoxia sustained in the interval between the time the microspheres were loaded into syringes and the time they exited the microcatheters may have contributed to loss of viability. Future studies in which electron microscopy is used to investigate the mechanical effects of transcatheter injection on ultrastructural cell architecture may advance our understanding of these processes.

Despite a decrease in the percentage of viable cells per microsphere, our findings did not demonstrate a substantial change in glucose metabolism following transcatheter injection. The difference in the viability percentage may not have been translated into a difference in metabolism in the early period (days 1–3) because of the limitations of our experimental technique. Since measurements of metabolic rate were averaged throughout a 3-day period, early changes (initial 24 hours) could not be detected. In addition, rapid initiation of cell growth following transcatheter injection may have further diminished our ability to detect early metabolic alterations. The absence of metabolic differences during the later period is expected, since the hydrogel matrix in a medium has a maximum capacity for cells, and small decreases in cell numbers are compensated by cell proliferation (31). This recovery phenomenon was probably not observed in our viability study, since we measured the percentage of viable cells per microsphere and not the absolute number of viable cells.

In conclusion, our findings demonstrate that 1% low-viscosity high-mannuronic alginate microspheres harboring fibroblast grafts can be delivered through a microcatheter without compromising the sequestration function of the hydrogel if the microsphere diameter does not exceed the catheter diameter. Transcatheter injection does cause a small but acceptable decrease in the percentage of viable cells per microsphere; however, the observed cell loss did not result in a measurable reduction of glucose metabolism in our study.

Practical applications: In this study, we have shown that microsphere-encapsulated cell preparations can be successfully delivered through a microcatheter without major alterations in microsphere structure, cell viability, or cell function. The ability to effectively deliver viable, metabolically active cell grafts through a microcatheter may enable the clinical treatment of diabetes mellitus and other disorders of cell function by means of minimally invasive endovascular cell transplantation. Clinical research has shown that nonencapsulated islet cell preparations delivered with percutaneous portal vein embolization through a 5-F catheter can survive in the liver and render patients with type 1 diabetes mellitus insulin independent (32). The use of microcatheters and hydrogel microspheres as embolic carrier agents for endovascular cell transplantation has many advantages, however. Microcatheters permit subselective delivery of cells to precise target vessel branches in specific anatomic locations. As discussed previously, the physical characteristics of hydrogel microspheres are well suited to the technical demands of transcatheter injection. The matrix of the microsphere also permits the application of tissue engineering technology to the design of highly specialized cell-carrier devices. Perhaps most important is that microspheres have the potential to serve as immunologic sanctuaries for transplanted cells and to eliminate the need for immunosuppression. Although this work demonstrates that the immunoisolation properties of alginate microspheres are preserved following microcatheter injection, additional work with animal models are needed to confirm this.


   ACKNOWLEDGMENTS
 
We are grateful to Steven Woodard, MS, for his technical support with our confocal microscopy work. We thank Linda Donoff for her assistance in preparation of the manuscript.


   FOOTNOTES
 
Author contributions: Guarantor of integrity of entire study, T.A.; study concepts and design, I.C., T.A.; literature research, T.A., H.J.C., A.S.; experimental studies, T.A., G.G.S.; data acquisition, T.A., G.G.S.; data analysis/interpretation, D.F.K., T.A., A.S., H.J.C.; statistical analysis, S.M.W.; manuscript preparation and definition of intellectual content, A.S., T.A., D.F.K., H.J.C.; manuscript editing, A.S., T.A., D.F.K., H.J.C., G.G.S.; manuscript revision/review, T.A., A.S., H.J.C., D.F.K., G.G.S., J.E.D.; manuscript final version approval, all authors.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
DISCUSSION
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