AJP - Heart Email Content Delivery
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 288: H584-H590, 2005. First published September 30, 2004; doi:10.1152/ajpheart.00690.2004
0363-6135/05 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
288/2/H584    most recent
00690.2004v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (12)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Kim, S.
Right arrow Articles by Johnson, P. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Kim, S.
Right arrow Articles by Johnson, P. C.

Aggregate formation of erythrocytes in postcapillary venules

Sangho Kim,1 Aleksander S. Popel,2 Marcos Intaglietta,1 and Paul C. Johnson1

1Department of Bioengineering, University of California, San Diego, La Jolla, California; and 2Department of Biomedical Engineering, Johns Hopkins University, Baltimore, Maryland

Submitted 13 July 2004 ; accepted in final form 17 September 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The purpose of the present study was to obtain information on erythrocyte aggregate formation in vivo. The movements of erythrocytes in postcapillary venules of the rat spinotrapezius muscle at various flow rates were recorded with a high-speed video camera before and after infusion of dextran 500. To distinguish aggregates, the following criteria were used: 1) a fixed distance (4 µm) between the center points of two adjacent cells, 2) lack of visible separation between the adjacent cells, and 3) movement of the adjacent cells in the same direction. Without dextran 500 infusion, 11 and 5% of erythrocytes formed aggregates in low (33.2 ± 28.3 s) and high pseudoshear (144.2 ± 58.3 s) conditions, respectively, based on the above criteria. After dextran 500 infusion, 53% of erythrocytes satisfied the criteria in the low pseudoshear condition (26.5 ± 17.0 s) and 13% of erythrocytes met the criteria in the high pseudoshear condition (240.0 ± 85.9 s), indicating erythrocyte aggregation is strongly associated with shear rate. Approximately 90% of aggregate formation occurred in a short time period (0.15–0.30 s after entering the venule) in a region 15 to 30 µm from the entrance. The time delay may reflect rheological entrance conditions in the venule.

hemodynamics; in vivo blood rheology; in vivo microscopy

RED BLOOD CELL AGGREGATION is a prominent feature in humans and other athletic species but not sedentary species (30). Numerous in vitro studies have shown that aggregation of blood increases as shear rate decreases. Aggregation also depends on hematocrit and the concentration of macromolecules in the plasma or suspending medium (7, 10, 15, 16, 19, 25, 32). However, the circumstances in which aggregation occurs in vivo and its possible importance in circulatory function are not well understood.

Previous whole organ studies in the dog and cat, in which red blood cell aggregation normally occurs, have shown that venous vascular resistance in skeletal muscle increases as arterial pressure and blood flow rate decrease and provided evidence that this effect is dependent on red blood cell aggregation (11, 23, 24, 35). From these studies, it was postulated that aggregate formation increased resistance to flow in the venular network. Recently, Bishop and co-workers (9) reported that, in rodents treated with high molecular mass dextran to induce aggregation, velocity profiles of blood flow in venules became blunted at low flow rates and red blood cells formed aggregates, which would explain, in part, the dependence of venous resistance on flow rate.

Current knowledge of the process of aggregate formation is based on in vitro studies using various systems including transparent cone-and-plate viscometers, glass capillary tubes, or narrow-gap flow chambers. These instruments make it possible to visualize the time course and degree of aggregation formation (1–4, 12, 14, 31, 33, 34). These investigations report a wide range of time scales (5–200 s) for aggregate formation, indicating that the rate of red blood cell aggregation might be associated with the dimensions and geometry of the experimental system. As a consequence, direct application of these findings to aggregate formation in venular networks is not feasible.

To our knowledge, there have been no studies on time course and process of aggregate formation in vivo. The purpose of the present study, therefore, was to provide such information. Postcapillary venules were selected for the study based on the assumption that aggregates would not be present in capillaries and this would be the first site in the venous network where aggregation could develop. The study was done using the spinotrapezius muscle preparation of the rat, an animal whose blood normally shows negligible red blood cell aggregation tendency (6). Intravenous infusion of dextran 500 was used to induce aggregation at levels seen in the human.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Experimental setup. An intravital microscope (Ortholux II, Leitz) transilluminated with a 100-W mercury lamp (model 1149, Walker Instruments, Scottsdale, AZ) was used with an Olympus x40 water-immersion objective and a long working distance condenser (Instec, Boulder, CO), which have numerical apertures of 0.7 and 0.35, respectively. Figure 1 shows the schematic diagram of the microscope system. A blue filter (transmittance peak 400 nm, model 59820, Spectra-Physics, Stratford, CT) was used to enhance the contrast between the red blood cells and the background field. Focused images viewed on a computer monitor were projected onto a high-speed video camera (model FASTCAM ultima SE, Photron USA) capable of recording up to 4,450 frames/s and storing up to 24,576 frames in the internal memory and was viewed on a computer monitor. This arrangement provided a full-frame field of 200 x 200-µm2 view. The high-speed video camera was connected to and controlled by a microcomputer (Intel Pentium IV). All the image files stored in the computer were transferred to a compact disk (CD-Writer Plus 7200e; Hewlett-Packard).



View larger version (26K):
[in this window]
[in a new window]
  		Fig. 1. Schematic diagram of the experimental setup. Details are described in Experimental setup. The high-speed video is capable of recording red blood cell movements in both capillaries and postcapillary venules at a wide range of flow rates.

 
Animal preparation. Eight male Wistar-Furth rats weighing 162 ± 25 g were used for the present study. Animal handling and care were provided in accordance with the procedures outlined in the Guide for the Care and Use of Laboratory Animals (National Institutes of Health, 1996). The study was approved by the local Animal Subjects Committee. The rats were anesthetized with an intraperitoneal injection of 50 mg/kg pentobarbital sodium (Abbott). Additional anesthetic was administered intravenously throughout the experiment as needed. The animal was placed on a heating pad to maintain body temperature of 37°C during surgery. A tracheal tube (ID = 1.57 mm, OD = 2.08 mm) was inserted to assist breathing, the jugular vein was catheterized for administration of anesthetic or dextran 500 during the course of the experiment, and the carotid artery was catheterized to measure arterial pressure and withdraw blood as needed for pressure reductions. All catheters were filled with a solution of heparinized saline (30 IU/ml) to prevent clotting. The rat spinotrapezius muscle was exteriorized and prepared for intravital microscopy as described in previous studies (8, 9). A temperature probe was placed beside the muscle so that temperature could be monitored and maintained at 37°C during the experiment by a heating element attached to the animal platform.

Pressure, hematocrit, and aggregation measurements. Arterial pressure was continuously recorded using a physiological data-acquisition system (MP 100 System, BIOPAC Systems, Goleta, CA). The pressure data were transferred and stored in a microcomputer during the experiment. Blood samples of ~0.1 ml were withdrawn from the carotid artery catheter during an experiment for hematocrit and degree of red blood cell aggregation measurements, which were taken during control as well as after infusion of Dextran 500. Hematocrit was determined after centrifugation with a microhematocrit centrifuge (Readacrit, Clay Adams). The degree of red blood cell aggregation was assessed from duplicate measurements on a 0.035-ml blood sample with a photometric rheoscope (Myrenne Aggregometer, Myrenne, Roentgen, Germany). The aggregation index (M) obtained in the present study was based on the 10-s setting.

Adjustment of red blood cell aggregability and hematocrit. Experiments were performed on animals both under normal (nonaggregating) conditions and after induction of red blood cell aggregation with dextran 500 (average molecular mass 460 kDa; Sigma) dissolved in saline (6%) and infused in 50-mg/kg increments over the course of 2–3 min (total 200 mg/kg body wt). This represents a plasma dextran concentration of ~0.6%. Hematocrit and aggregation index values were determined 15 min after dextran 500 infusion. There was no discernable adverse reaction (e.g., visible swelling of the limbs) to the dextran 500 infusion in any of the rats used in these investigations.

To obtain clear and distinct images of both individual red blood cells and aggregates in the venule, it was necessary to reduce local hematocrit. This was achieved by reduction of systemic hematocrit with infusion of autologous plasma obtained from a donor animal. The blood volume was maintained by exchanging the same amount of blood with plasma (1–2 ml). Mean arterial pressure was monitored during and after blood withdrawal and plasma infusion until a steady-state value, which was not significantly different from the control arterial pressure, was reestablished.

To estimate the tube hematocrit in the postcapillary venules, symmetric cylindrical vessel geometry was assumed. The dimensions of the postcapillary venule, the mean red blood cell volume of the rat (~52 µm3) (5), and the number of red blood cells present in the venule at a specific moment were used for the estimation.

Experimental protocol. Before the experiment began, a blood sample was taken from the carotid artery to obtain control values of hematocrit and aggregation index. Skeletal muscle postcapillary venules with two to three supply capillaries in the field of view were selected for the study on the basis of the criteria of stable flow as well as clear focus and contrast of the image. The microscope was focused on the equatorial plane of the venule, and the high-speed video camera was used to record the movements of red blood cells in both the capillaries and venule for ~30 s. The frame rate was adjusted to maximize image clarity for the specific flow and background conditions. The frame rates of 500–2,250 frames/s were used in this study. In general, higher frame rates gave us more distinct edge definition of red cells but lower contrast with background. Thus the frame rates of 500–750 frames/s were used for reduced arterial pressures and higher frame rates were used for normal arterial pressures.

The microcirculatory observations were repeated at reduced flow rates. For this purpose, blood was removed via the carotid artery into a heparinized syringe. An average of 1.0 ± 0.5 ml of blood was withdrawn at a rate of ~2 ml/min. After the reduced pressure stabilized and steady blood flow was reestablished, the video image of the vessels was recorded for ~1 min. The blood withdrawn was reinfused after the recording, and in all cases, the arterial pressure was monitored until it returned to a stable value. After completion of the above procedures, red blood cell aggregation was induced with infusion of dextran 500 and the experimental protocol described above was repeated. In some instances, it was necessary to find another suitable venule due to either unsteady flow or cessation of flow in one or more of the supply capillaries of the venule used in the control study.

Determination of red blood cell and wall positions. Digital video clips stored in the control unit of the high-speed camera were transferred to a hard drive in the microcomputer for analysis and storage. Using Adobe Premier 5.1, the video files were then converted to image files (either TIFF or BMP format). Image magnification was determined from the recorded image of a stage micrometer under transillumination. Figure 2 shows a videomicrograph of a typical vessel studied under transillumination (top) and a schematic diagram of the vessel (bottom), respectively.



View larger version (73K):
[in this window]
[in a new window]
  		Fig. 2. Videomicrograph and schematic diagram of a representative postcapillary venule with 2 supply capillaries. A blue filter was used to enhance the contrast between the red blood cells and the background field. The microscopic system provided a full-frame field of 200 x 200 µm2. The postcapillary venule image shown here was a portion of the field. The contrast of image was increased for publication.

 
From an image such as shown in Fig. 2, top, x-y coordinate data for each red blood cell image were obtained using an image analysis software package (SigmaScan Pro 4.0, SPSS). Depending on the flow velocity and framing rate, every fifth to fiftieth frame was chosen for the determination of instantaneous positions of each red blood cell coming from capillaries and passing through the postcapillary venule. A total of 12 postcapillary venules were used in the present study and for each venule, 24–30 red blood cells were selected and analyzed. Similarly, venular wall position was determined from the transillumination image. However, in the latter case, an averaged value from five measurements was used to minimize human measurement error. Error associated with marking the center of red blood cell images on the image analysis software was estimated to be ±0.5 µm based on 10 repeated measures on a group of 5 cells.

The mean velocity of each red blood cell was obtained from the position data of the cells and time interval between two position measurements. The velocities were averaged and used for the mean velocity (V) of blood flow in the venule. Pseudoshear rate in the venule was calculated from the following equation

(1)

where {gamma} is pseudoshear rate and D is average diameter. The average diameter was determined from five measurements at different locations along the venule using the image analysis software.

Criteria for red blood cell aggregate formation. To determine whether adjacent red blood cells form an aggregate, the following criteria were used: 1) a fixed distance (4 µm) between the center points of two adjacent cells based on measurements in stationary aggregates in vitro, 2) lack of visible separation between the adjacent cells, and 3) movement of the two adjacent cells in the same direction.

To establish the first criterion, a blood sample was removed from a dextran-treated animal via the carotid artery into a heparinized syringe and a drop was placed on a glass slide and covered with a glass coverslip. The field containing the in vitro red blood cell aggregates was recorded for ~10 s and converted into image files. The distance between centers of two adjacent red blood cells in selected aggregates was measured using image analysis software. The mean distance between centers of two unstressed red blood cells obtained from the in vitro experiment was 3.4 ± 0.1 µm. Bishop and co-workers (8) reported that the minimum width of red blood cells in 50-µm skeletal muscle venules, representing cell thickness, was ~3.7 µm. Considering the error of human measurement for the cell location in our study, which was ±0.5 µm, a value of 4 µm or less between centers of two cells was used as the criterion for aggregate formation.

Shown in Fig. 3 is a schematic diagram of the different cell configurations for each case (nonaggregating cells, aggregate, or possible aggregate) we observed in vivo. The distance between the centers of adjacent cells, as shown in Fig. 3, was directly calculated from the position data of the cells passing through the postcapillary venule in the field of view. The aggregate case (see Fig. 3C) consisted of either two or more cells in parallel with the flow direction or crescent-shaped cells in tandem. In the case of possible aggregate, a few unusual cell configurations with two crescent-like or/and oval-like cells (see Fig. 3B) were observed, whereas the configurations designated as possible aggregates showed similar shapes but larger center-to-center distances (>4 µm) compared with apparent aggregates. If there was distinct separation between two adjacent red blood cells as shown in Fig. 3A, the cells were classified as nonaggregating cells.



View larger version (10K):
[in this window]
[in a new window]
  		Fig. 3. Typical shapes of red blood cells observed in the study for 3 cases. A: nonaggregating cells. B: possible aggregates. C: aggregates. The arrows shown between 2 red cells indicate the center-center distance between the cells.

 
Statistical analysis. A paired t-test was used to determine differences in experimental and physiological parameters between normal and dextran-treated animals. The degree of aggregation was compared in four groups as follows: normal arterial pressure, normal arterial pressure with dextran 500 infusion, reduced flow condition, and reduced flow condition with dextran 500 infusion. At least two animals were used for each condition that provided the experimental data from three different venules. All of the data are reported as means ± SD. The statistical significance of differences in the aggregate formation of red blood cells was determined by comparing the percentage of aggregates in each case, using a one-way ANOVA test with subsequent Scheffé's, Fisher's least significant difference, and Bonferroni tests. All statistical tests were done using a commercially available software Package (SPSS 11.0 for Windows). For all tests, P < 0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Hematocrit, degree of aggregation, and arterial pressure. As noted in MATERIALS AND METHODS, for better observation of aggregation formation, normal systemic hematocrit (42.4 ± 2.1%) was reduced by hemodilution with plasma. The reduced hematocrit in the control period (27.1 ± 2.4%) was not significantly different from that after dextran treatment (27.7 ± 3.2%, P > 0.05). The mean diameter and length of postcapillary venules selected in the present study were 12.0 ± 1.4 and 83.1 ± 35.7 µm, respectively. Based on these values, the tube hematocrit was 9.4 ± 2.1% for the 12 postcapillary venules used in the study. The thickness of the endothelial glycocalyx was not considered in this calculation but its possible effect is considered in DISCUSSION.

Under control conditions for normal rats, the index of aggregation, M, was 0.0 in all cases, and the mean arterial pressure was 113.3 ± 5.8 mmHg. The reduced mean arterial pressure for normal rats was 56.3 ± 17.4 mmHg. There were no significant differences (P > 0.05) between the mean arterial pressures of normal and dextran-treated animals during the control and reduced flow situations. In dextran-treated rats, the index of aggregation was 16.9 ± 3.9 and the arterial pressures were 110.0 ± 8.7 and 55.7 ± 9.0 mmHg during control and reduced flow situations, respectively. No significant difference (P > 0.05) in the degree of aggregation was found between normal and reduced flow conditions in dextran-treated rats.

Time course of red blood cell aggregate formation. The time course of red blood cell aggregate formation in a postcapillary venule is shown in Fig. 4. A sample group of eight red blood cells in two capillaries at the entrance region of the venule was identified (see Fig. 4A) and monitored over time. Once the individual cells entered the venule, as shown in Fig. 4, B–D, the separation between red blood cells tended to decrease and some cells formed aggregates. After red blood cells formed aggregates, the adherent cells moved together in the same direction. The distance between the red blood cells that formed an aggregate was monitored until the aggregate moved out of focus, which was usually ~80 µm from the entrance.



View larger version (135K):
[in this window]
[in a new window]
  		Fig. 4. Time history of red blood cell aggregate formation in a postcapillary venule. Representative videomicrographs of 13.7-µm postcapillary venule at t = 0.00 s (A), t = 0.10 s (B), t = 0.20 s (C), and t = 0.32 s (D) after dextran treatment at a pseudoshear rate of ~10 s. The dotted lines indicate the walls of blood vessels. The red cell images shown here were also enhanced.

 
As shown in Fig. 4, the boundaries of individual red blood cells were distinct before they formed aggregates but became indistinct in the region between adjoining cells once aggregates formed. If the adjacent cells maintained a distance <4 µm and moved in the same direction with no distinct separation in between, the two (or more) cells were defined as an aggregate.

Distance between the centers of two adjacent red blood cells in an aggregate. A total of 306 red blood cells were analyzed. Between 72 and 78 red blood cells were investigated for each of four conditions: nondextran condition at control and reduced arterial pressure and dextran treatment at control and reduced arterial pressure. Figure 5 shows the distance changes between the centers of two adjacent red blood cells over time for three cases: aggregate, possible aggregate, and nonaggregating cells. Note that the distance measurement covers the period in which the red blood cells were flowing in capillaries as well as in a postcapillary venule as shown in Fig. 5. The distance data indicate that in two cases the red blood cells came from two different capillaries and in one case from the same capillary.



View larger version (27K):
[in this window]
[in a new window]
  		Fig. 5. Distance changes between the centers of 2 adjacent red blood cells over time for 3 cases: aggregate, possible aggregate, and nonaggregating cells. Data points in the gray area represent red blood cells in capillaries, whereas other data points are from red blood cells in a postcapillary venule. Dotted line indicates 4 µm, one of criteria used in the present study.

 
For the case in which an aggregate formed, the distance between centers of the two cells entering from two supply capillaries decreased gradually and eventually stabilized below 4 µm in the postcapillary venule. The two cells tumbled after aggregate formation and the center-to-center distance varied between ~2 and 4 µm. In the possible aggregate case, the distance fluctuated around 4 µm but did not stay below this value while the other criteria were met. For the case of nonaggregating cells, the distance between the cell centers was always larger than 4 µm.

Red blood cell aggregation in postcapillary venules. As shown in Fig. 6, without dextran 500 infusion, 11 and 5% of red blood cells formed aggregates in low ({gamma} = 33.2 ± 28.3 s) and high pseudoshear ({gamma} = 144.2 ± 58.3 s) conditions, respectively, based on the criteria. With dextran 500 infusion, 53% of red cells satisfied the criteria in the low pseudoshear condition ({gamma} = 26.5 ± 17.0 s), whereas 13% of red cells met the criteria in the high psuedoshear condition ({gamma} = 240.0 ± 85.9 s), indicating red cell aggregation is highly associated with shear rate. The percentages of aggregating and nonaggregating cells with dextran treatment at reduced arterial pressure were significantly different (P < 0.01) from those for any of the other three conditions. The latter were not significantly different (P > 0.05) from each other.



View larger version (24K):
[in this window]
[in a new window]
  		Fig. 6. Experimental results of red blood cell aggregate formation in postcapillary venules based on the 3 criteria. Values are means ± SD; n = 3 in each group. *P < 0.01.

 
Rate of red blood cell aggregate formation. Figure 7 shows red blood cell aggregate formation as a function of time and position in postcapillary venules. In this analysis, the aggregation data used were only for dextran treatment at reduced arterial pressure. There was no red blood cell aggregate formation at the immediate entrance region of postcapillary venules. Aggregate formation began rather suddenly at ~30 ms elapsed time when the distance of the red blood cell from the entrance was 15 µm and then increased rapidly as the elapsed time increased to ~100 ms and the distance traveled increased to 22 µm. In fact, ~90% of the 17 aggregates we observed were formed in the region between 15 and 30 µm of the entrance to the postcapillary venule and it took less than 0.3 s to form all aggregates. In the remaining 30-µm segment we were able to visualize, the rate of aggregate formation was very low despite the fact that ~50% of the red blood cells were still in the nonaggregated state.



View larger version (20K):
[in this window]
[in a new window]
  		Fig. 7. Aggregate formation of red blood cells as a function of time and position in postcapillary venules (11.2 ± 2.3 µm). {bullet} Indicate the percentage of red cells in all aggregates, whereas {triangleup} represent the percentage of larger aggregates (3–5 cells).

 
The open triangle symbols in Fig. 7 represent the larger aggregates, showing that ~50% of total aggregates were formed by three or more red blood cells. In general, aggregates were first formed by two cells joining. As those binary aggregates moved downstream, six became larger by individual red blood cells or aggregates joining the binary aggregate. In two instances, two binary aggregates coupled to each other, and in one instance, three cells combined simultaneously to form one large aggregate. The largest aggregate observed in this study consisted of five red blood cells. Rotary motion (tumbling) was observed in both aggregates and individual red cells. Although the speed of such motion could not be quantified, we observed rotation in 12% of aggregates and 14% of individual cells in the ~70-µm length of the venule we observed.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The purpose of the present study was to provide information on the time course and process of red blood cell aggregate formation in vivo. To our knowledge, this is the first such study. Postcapillary venules were chosen for the study because this is the first site in the venous network where aggregation could develop. No red blood cell aggregates were observed in supply capillaries. We show that most of the aggregate formation occurs in a region 15–30 µm from the entrance to postcapillary venules. It appears that a much shorter time period is required for red blood cells to form aggregates in vivo compared with those commonly reported for in vitro studies (5–200 s) (1–4, 12, 14, 31, 33, 34).

The major source of uncertainty in distinguishing aggregates based on the criteria presented in MATERIALS AND METHODS was in determining the center-to-center distance of 4 µm. The distance measurement was based on the horizontal plane coordinate, although actual distance should include the z coordinate as well. Therefore, this measurement might underestimate the actual separation between two red blood cells at different levels moving with the same velocity in a postcapillary venule. To minimize error due to overlap of the erythrocyte images, hematocrit was reduced to a level at which the images of red blood cells were distinct.

The degree of red cell aggregability induced by high molecular mass polymers is dependent on hematocrit. It has been pointed out that the endothelial glycocalyx, which is not observable with light microscopy, could alter local hematocit and change flow patterns in small vessels such as postcapillary venules and capillaries (17, 28). In the present study, assuming a 0.5-µm-thick glycocalyx in the venule as suggested by Damiano and co-workers (17, 26), the tube hematocrit would be 11.3 ± 2.6%, a 20% increase compared with the estimated value of 9.4 ± 2.1% without a glycocalyx.

It is well known that red blood cell aggregates are formed in the presence of long-chain macromolecules such as dextran 500. In vitro studies on the formation of aggregates show a strong relationship between aggregate size and shear rate (4, 8, 22). Goldsmith and Marlow (22) reported that, in suspensions of <2% hematocrit, only single cells were observed at shear rates above ~50 s, whereas large aggregates with 10–20 red blood cells were seen at shear rates below ~4 s. More recently, Yedgar and co-workers (12) found that red blood cells (hematocrit = 10%) formed small aggregates with three to four cells at a shear rate of ~70 s. Bishop et al. (8) found that aggregates were two to three times wider than the diameter of a single red blood cell at the low pseudoshear rates tested, less than ~10 s, whereas at pseudoshear rates above ~50 s the average width of individual particles was not significantly different from the red blood cell diameter. These previous results agree with the results shown in Fig. 5, supporting the criteria for aggregate formation.

The time required to begin aggregate formation reported from in vitro studies is longer than we report and varies with the system used. In previous studies, red blood cell aggregation was observed following a sudden change in shear rate, which might require a longer time period for flow stabilization (21, 36), compared with steady flows at low shears in the present study. Alonso and co-workers (1–3) found that cell-free plasma spaces resulting from red cell aggregation started to develop 5 to 10 s after sudden reduction of flow rate in straight capillary tubes (25- to 100-µm ID). Chen and co-workers (12, 14) observed no significant change of particle sizes in human blood in the first 10–20 s after stopping flow in a 40-µm gap flow chamber (hematocrit = ~10%) and a sudden development of aggregation after that.

In a study of whole blood viscosity with a Contraves viscometer, red cell aggregates in human blood (hematocrit = 47%) appeared to form in ~2 s with shear rate reduction to 0.7 s (13). However, it may take less time to form binary aggregates like those observed in our study because the shear rate of 0.7 s used in that study was low enough to form large three-dimensional aggregates. Hematocrit is also important. Shiga and co-workers (33, 34) found that the development of one-dimensional short rouleaux (composed of 2 or 3 red blood cells) required longer than 150 s at a steady shear rate of 7.5 s in the transparent cone-and-plate viscometer with a hematocrit of 0.25%. The much shorter time required for red blood cell aggregation to begin in the present study (30 ms) and reach completion (0.3 s) likely reflects the higher hematocrit and/or smaller scale of the system than in previous in vitro studies.

An important finding of our study, as shown in Fig. 7, is the lack of red blood cell aggregate formation at the entry region 15 µm from the branching point of postcapillary venules. Das and co-workers (18) performed a numerical study of blood flows in small venular bifurcations and showed that the velocity profiles became fully developed ~1.5 diameters downstream from the bifurcation. Therefore, it appears that the absence of aggregation in the entrance region reflects the distance required for convergence of the streamlines that brings the red blood cells close together.

Another significant finding is that aggregation was essentially complete 30 µm from the branch point, although 47% of the cells had not formed aggregates. Because the tube hematocrit for the venules used in this study was relatively low (9–11% depending on assumed glycocalyx width), not all cells would come in contact with another cell. The age of the red blood cells could be involved. Meiselman (27) demonstrated that relatively young red blood cells had approximately three times lower aggregability compared with older cells.

A few aggregates (5 and 11%, respectively, of total cells) according to our criteria were formed in postcapillary venules of normal rats at control and reduced shear rates, although the aggregability (M) of the blood measured with the Myrenne Aggregometer was zero. It is possible that these aggregates represent short-lived red cell clusters brought in close contact solely by hemodynamic forces as suggested by Osterloh et al. (29). They reported a mean particle size considerably greater than that of red cells at high shear rates in pre- and postcapillary vessels (~25 µm) using a spatial Fourier analysis of light intensity patterns. These clusters are not linked together by mechanisms engendered by macromolecules.

Approximately 13% of cells met the criteria for aggregation at the high pseudoshear rate after dextran treatment. Not surprisingly, these aggregates formed further downstream, between 40 and 55 µm from the entrance to the postcapillary venule. Osterloh and coworkers (29) found that the pattern size of particles in pre- and postcapillary vessels increased at pseudoshear rates both higher and lower than ~150 s after infusion of dextran 250 (800 mg/kg body wt). The presence of aggregation at high pseudoshear rates in their study and ours may be explained not only by the hemodynamic forces mentioned above but also by the macromolecules of dextran once the hemodynamic forces bring red blood cells into close contact.

Previous studies have shown that the formation of red cell aggregates in the venules contributes significantly to the pressure drop in that bed, thus influencing the overall energy dissipation in the venous circulation (9, 11). In addition to the effect of increased particle size on velocity profiles, the tumbling of aggregates disturbs the flow and leads to the dissipation of energy, resulting in increased blood viscosity at low shear rates (20). We observed in this study that a few (12%) of the aggregates tumbled. However, the distance that the aggregates could be observed in the postcapillary venules after aggregate formation was short, and this might have contributed to the low percentage of tumbling aggregates observed. It appeared that the tumbling motion of aggregates or individual red cells was very slow compared with their movements in a flow direction.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grants HL-52684, HL-62354, and HL-64395.


   ACKNOWLEDGMENTS
 
The authors thank Dr. P. Cabrales, K. Flores, S. Dunning, J. H. Park, and R. Pang for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. C. Johnson, Dept. of Bioengineering, Univ. of California, San Diego, La Jolla, CA 92093-0412 (E-mail: pjohnson{at}bioeng.ucsd.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
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Alonso C, Pries AR, and Gaehtgens P. Time-dependent rheological behaviour of blood flow at low shear in narrow horizontal tubes. Biorheology 26: 229–245, 1989.[ISI][Medline]
  2. Alonso C, Prices AR, and Gaehtgens P. Time-dependent rheological behavior of blood at low shear in narrow vertical tubes. Am J Physiol Heart Circ Physiol 265: H553–H561, 1993.[Abstract/Free Full Text]
  3. Alonso C, Prices AR, Kiesslich O, Lerche D, and Gaehtgens P. Transient rheological behavior of blood in low-shear tube flow: velocity profiles and effective viscosity. Am J Physiol Heart Circ Physiol 268: H25–H32, 1995.[Abstract/Free Full Text]
  4. Barshtein G, Wajnblum D, and Yedgar S. Kinetics of linear rouleaux formation studied by visual monitoring of red cell dynamic organization. Biophys J 78: 2470–2474, 2000.[Abstract/Free Full Text]
  5. Baskurt OK. Deformability of red blood cells from different species studied by resistive pulse shape analysis technique. Biorheology 33: 169–179, 1996.[CrossRef][ISI][Medline]
  6. Baskurt OK, Farley RA, and Meiselman HJ. Erythrocyte aggregation tendency and cellular properties in horse, human, and rat: a comparative study. Am J Physiol Heart Circ Physiol 273: H2604–H2612, 1997.[Abstract/Free Full Text]
  7. Baskurt OK, Meilselman HJ, and Kayar E. Measurement of red blood cell aggregation in a "plate-plate" shearing system by analysis of light transmission. Clin Hemorheol Microcirc 19: 307–314, 1998.[ISI][Medline]
  8. Bishop JJ, Nance PR, Popel AS, Intaglietta M, and Johnson PC. Relationship between erythrocyte aggregate size and flow rate in skeletal muscle venule. Am J Physiol Heart Circ Physiol 286: H113–H120, 2004.[Abstract/Free Full Text]
  9. Bishop JJ, Nance PR, Popel AS, Intaglietta M, and Johnson PC. Effect of erythrocyte aggregation on velocity profiles in venules. Am J Physiol Heart Circ Physiol 280: H222–H236, 2001.[Abstract/Free Full Text]
  10. Boynard M and Lelievre JC. Size determination of red blood cell aggregates induced by dextran using ultrasound backscattering phenomenon. Biorheology 27: 39–46, 1990.[ISI][Medline]
  11. Cabel M, Meiselman HJ, Popel AS, and Johnson PC. Contribution of red blood cell aggregation to venous vascular resistance in skeletal muscle. Am J Physiol Heart Circ Physiol 272: H1020–H1032, 1997.[Abstract/Free Full Text]
  12. Chen S, Barshtein G, Gavish B, Mahler Y, and Yedgar S. Monitoring of red blood cell aggregability in a flow-chamber by computerized image analysis. Clin Hemorheol 14: 497–508, 1994.
  13. Chen S, Dormandy J, Ernst E, and Matrai A. Clinical Hemorheology: Biophysics. Hingham, MA: Kluwer Academic, 1987, chapt. 2.
  14. Chen S, Gavish B, Zhang S, Mahler Y, and Yedgar S. Monitoring of erythrocyte aggregate morphology under flow by computerized image analysis. Biorheology 32: 487–496, 1995.[CrossRef][ISI][Medline]
  15. Cloutier G, Qin Z, Durand L, and The BG. Power Doppler ultrasound evaluation of the shear rate and shear stress dependences of red blood cell aggregation. IEEE Trans Biomed Eng 43: 441–450, 1996.[CrossRef][ISI][Medline]
  16. Cloutier G and Shung KK. Study of red cell aggregation in pulsatile flow from ultrasonic Doppler power measurements. Biorheology 30: 443–461, 1993.[ISI][Medline]
  17. Damiano ER. The effect of the endothelial-cell glycocalyx on the motion of red blood cells through capillaries. Microvasc Res 55: 77–91, 1998.[CrossRef][ISI][Medline]
  18. Das B, Enden G, and Popel AS. Stratified multiphase model for blood flow in a venular bifurcation. Ann Biomed Eng 25: 135–153, 1997.[ISI][Medline]
  19. Fontaine I, Savery D, and Cloutier G. Simulation of ultrasound backscattering by red cell aggregates: effect of shear rate and anisotropy. Biophys J 82: 1696–1710, 2002.[Abstract/Free Full Text]
  20. Fung YC. Biomechanics: Mechanical Properties of Living Tissues, 2nd Edition. New York: Springer, 1993.
  21. Gaspar-Rosas A and Thurston GB. Erythrocyte aggregate rheology by transmitted and reflected light. Biorheology 25: 471–487, 1988.[ISI][Medline]
  22. Goldsmith HL and Marlow J. Flow behavior of erythrocytes. I. Rotation and deformation in dilute suspensions. Proc R Soc Lond B Biol Sci 182: 351–384, 1972.
  23. House SD and Johnson PC. Microvascular pressure in venules of skeletal muscle during arterial pressure reduction. Am J Physiol Heart Circ Physiol 250: H838–H845, 1986.[Abstract/Free Full Text]
  24. Johnson PC and Hanson KM. Effect of arterial pressure on arterial and venous resistance of intestine. J Appl Physiol 17: 503–508, 1962.[Abstract/Free Full Text]
  25. Kim SY, Miller IF, Sigel B, Consigny PM, and Justin J. Ultrasonic evaluation of erythrocyte aggregation dynamics. Biorheology 26: 723–736, 1989.[ISI][Medline]
  26. Long DS, Smith ML, Pries AR, Ley K, and Damiano ER. Microviscometry reveals reduced blood viscosity and altered shear rate and shear stress profiles in microvessels after hemodilution. Proc Natl Acad Sci USA 101: 10060–10065, 2004.[Abstract/Free Full Text]
  27. Meiselman HJ. Red blood cell role in RBC aggregation: 1963–1993 and beyond. Clin Hemorheol 13: 575–592, 1993.
  28. Mulivor AW and Lipowsky HH. Role of glycocalyx in leukocyte-endothelial cell adhesion. Am J Physiol Heart Circ Physiol 283: H1282–H1291, 2002.[Abstract/Free Full Text]
  29. Osterloh K, Gaehtgens P, and Pries AR. Determination of microvascular flow pattern formation in vivo. Am J Physiol Heart Circ Physiol 278: H1142–H1152, 2000.[Abstract/Free Full Text]
  30. Popel AS, Johnson PC, Kameneva MV, and Wild MA. Capacity for red blood cell aggregation is higher in athletic mammalian species than sedentary species. J Appl Physiol 77: 1790–1794, 1994.[Abstract/Free Full Text]
  31. Schmid-Schönbein H, Gaehtgens P, and Hirsch H. On the shear rate dependence of red cell aggregation in vitro. J Clin Invest 47: 1447–1454, 1968.[ISI][Medline]
  32. Sennaoui A, Boynard M, and Pautou C. Characterization of red blood cell aggregate formation using an analytical model of the ultrasound backscattering coefficient. IEEE Trans Biomed Eng 44: 585–591, 1997.[CrossRef][ISI][Medline]
  33. Shiga T, Imaizumi K, Harada N, and Sekiya M. Kinetics of rouleaux formation using TV image analyzer. I. Human erythrocytes. Am J Physiol Heart Circ Physiol 245: H252–H258, 1983.[Abstract/Free Full Text]
  34. Shiga T, Imaizumi K, Maeda N, and Kon K. Kinetics of rouleaux formation using TV image analyzer. II. Rat erythrocytes. Am J Physiol Heart Circ Physiol 245: H259–H264, 1983.[Abstract/Free Full Text]
  35. Thulesius O and Johnson PC. Pre- and postcapillary resistance in skeletal muscle. Am J Physiol 210: 869–872, 1966.[Free Full Text]
  36. Thurston GB. Advances in Hemodynamics and Hemorheology: Viscoelastic Properties of Blood and Blood Analogs. London: Jai, 1996, chapt. 1.



This article has been cited by other articles:


Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
S. Kim, R. L. Kong, A. S. Popel, M. Intaglietta, and P. C. Johnson
Temporal and spatial variations of cell-free layer width in arterioles
Am J Physiol Heart Circ Physiol, September 1, 2007; 293(3): H1526 - H1535.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
S. Kim, J. Zhen, A. S. Popel, M. Intaglietta, and P. C. Johnson
Contributions of collision rate and collision efficiency to erythrocyte aggregation in postcapillary venules at low flow rates
Am J Physiol Heart Circ Physiol, September 1, 2007; 293(3): H1947 - H1954.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Heart Circ. Physiol.Home page
S. Kim, A. S. Popel, M. Intaglietta, and P. C. Johnson
Effect of erythrocyte aggregation at normal human levels on functional capillary density in rat spinotrapezius muscle
Am J Physiol Heart Circ Physiol, March 1, 2006; 290(3): H941 - H947.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
288/2/H584    most recent
00690.2004v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (12)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Kim, S.
Right arrow Articles by Johnson, P. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Kim, S.
Right arrow Articles by Johnson, P. C.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2005 by the American Physiological Society.