British Journal ofHaematology, 1977, 35, 71.

Myosin in Cultured Human Endothelial Cells

ANNE MOORE, ERIC A. JAFFE, CARL G. BECKER AND RALPH L. NACHMAN

Division of Haematology, Department of Medicine and Department o f Pathology, Corne11 University Medical College, N e w York

(Received I ]une 1976; accepted for publication ~ o ] u n e 1976)

SUMMARY. Myosin was isolated from cultured human endothelial cells by extrac- tion with 0.6 M KC1 and chromatography on Sepharose 4B. The extracted en- dothelial cell protein was identified as myosin by the characteristic ATPase profile, that is, the ATPase was activated by Ca2 + and EDTA and inhibited by MgZ +. On sodium dodecyl sulphate polyacrylamide gel el'ectrophoresis, the endothelial ccll myosin heavy chain migrated with a molecular weight of 200 ooo as did rabbit uterine and human platelet myosin heavy chains. A crude preparation of the cndothelial cell myosin reacted immunologically with an antiserum to platelet myosin, a smooth muscle type of myosin. In indirect immunofluorescence studies, antiserum to the purified endothelial cell myosin stained cultured endothelial cells in a fibrillar pattern. The fibrillar pattern was more intense when the endothelial cells were stained with antiserum to platelet myosin. The presence of myosin in the endothelial cell provides a basis for the contractility of these cells. This con- tractile property may plan an important role in the physiologic function of these cells.

Contractile proteins have been found in many non-muscle cells and may be important for cell motility, cell division, secretion, and phagocytosis (Pollard & Weihing, 1974; Adelstein, 1975). Human endothelial cells both in vivo and in vitro have been shown by indirect methods to contain contractile proteins which are antigenically similar to platelet and uterine actomyo- sin (Becker & Nachman, 1973 ; Jaffe et a!, 1973). Endothelial cell contractility might play a role in such physiologic processes as the regulation of vascular permeability, the inflam- matory response and the initiation of thrombosis. In order to study the contractile mechan- isms of these cells, we have isolated and characterized endothelial cell myosin. The purification of this protein was made possible by the recent development of two techniques. First, pure human endothelial cells free of contaminating smooth muscle cells or fibroblasts have become available in culture (Jaffe et af, 1973). Second, a method for extracting myosin from a small amount of cell protein has been developed (Ostlund et al, 1974).

I n this report we describe the partial purification and characterization of myosin from cultured human umbilical vein endothelial cells.

Correspondence: Dr Anne Moore, Division of Hematology, Department of Medicine, Cornell University Medical College, $25 East 68th Street, New York, N.Y. 10021, U.S.A.

72 A n n e Moore et al

MATERIALS AND METHODS

Endothelid cell culture. Endothelial cells were obtained from human umbilical cord veins and cultured as previously described (Jaffe et al , 1973).

Extraction of endothelial cell myosin. The method described by Ostlund et a! (1974) was used. All procedures were performed at 0-4"C. Endothelial cells grown to confluence were washed three times with phosphate buffered saline (0.145 M NaCI, 0.01 M phosphate buffer, pH 7.4) containing 2 mM dithiothreitol (DTT) (Sigma Chemical Co., St Louis, Missouri) and once with 0.15 M NaCl-2 I I ~ M DTT containing 0.5% bovine serum albumin (Miles Laboratories, Inc., Kankakee, Illinois) added to facilitate the uptake of the cells. The cells were removed with a rubber policeman and sedimented by centrifugation at 700 g for 15 min. The cell pellet was washed twice in 0.15 M NaC1-2 mM DTT by centrifugation at 700 g for 5 min and then frozen at - 70°C. The cell pellet was used usually within 2 or 3 weeks but equivalent results were found with pellets frozen up to 5 months. In a typical experiment, cells from approximately 40 Petri dishes ( IOO x 20 mm) or the equivalent number of cells from T-75 flasks or 2-litre roller bottles were used. This was equivalent to 30-40 mg of total cell protein.

To extract myosin, the cell pellets were thawed, resuspended in buffer A (0.6 M KCl, 1 5 mM Tris-HC1, pH 7.5 containing I-butanol (3%, v/v) and 2 mM DTT) and stirred inter- mittently for I h. The suspension was then centrifuged at 12 ooog for 10 min. The cell lysate supernatant, concentrated to I ml in dialysis tubing packed in dry Sephadex G-zoo gel (Pharmacia Fine Chemicals, Inc., Piscataway, N.J.), was applied to a column (0.9 x 60 cni) of Sepharose 4B (Pharmacia) which had been washed with buffer B (buffer A without butanol). The specimen was eluted with buffer B at a rate of 3 ml/h and 4 ml fractions were collected and analysed within 12 h for ATPase activity. In some experiments, an aliquot of the cell lysate supernatant was dialysed against buffer B for 24-38 h and then tested for ATPase activity in an attempt to estimate the recovery of ATPase activity in the column fractions.

Extraction ofplatelet myosin. Outdated platelets (3-10 d old), were kindly supplied by The New York Blood Center. The platelets were pooled, centrifuged at 300 g at room tempera- ture, washed twice with 0.9% NaCl, 0.3% Na citrate, pH 7.2, and stored at - 70°C for up to 3 weeks. Myosin was extracted from 50 g of thawed platelets by the method of Pollard et al

Rabbit crterine actomyosin. Rabbit uterine actomyosin was prepared as described previously (Becker & Nachman, 1973).

A T P a s e assay. Myosin ATPase is active in the presence of potassium and either calcium or EDTA but is inactive in the presence of magnesium (Seidel, 1969). In order to define a myosin-like ATPase activity in the endothelial cell preparation, the assay describcd by Adelstein et al (1971) was used. A 0.25-0.5 ml aliquot of the chromatography fraction was incubated at 37°C for 30 niin in 2 ml containing 2 mM ATP (Sigma Chemical Co.), 10 mM imidazole-HC1, pH 7 (J. T. Baker Chemical Co., Phillipsburg, NJ.), 0.6 M KCl, and either 2 mM EDTA, 10 n m CaCl, or 5 niM MgCl,. The ATP solutions and the column samples were incubated alone as controls. An inorganic phosphorus standard was included in each assay, The amount of inorganic phosphorus at the beginning and at the end of the

(1974).

Myosin in Endothelial Cells 73 incubation period was measured by a modification of the technique of Martin & Doty (1949). The results are expressed as ATPase units where I unit is that amount of enzyme that released I pmole of inorganic phosphorus per minute.

Protcin determination. All samples for protein determination were diluted with 0.15 M NaCl, precipitated with a final concentration of 10% trichloroacetic acid, and the precipitate solubilized with I N NaOH for I h. The protein was measured by the method of Lowry using bovine serum albumin as a standard (Lowry et al, 1951).

Immunization. Antibody to endothelial cell myosin was prepared by injecting a rabbit with a total of 350 pg of protein which had been dialysed against water and lyophilized. The protein was reconstituted with 0.9% NaCl, mixed with an equal volume of complete Freund’s adjuvant, and injected into footpads on two occasions 2 weeks apart. Two weeks later, the animal was boosted with an intramuscular injection of the antigen and this was repeated I month later. The antiserum was harvested 2 weeks after the last injection. Anti- body to platelet myosin was prepared in a similar manner using a total of 360 pg of protein.

Immunodifusion tests. These were performed in gels composed of 1.5% Agarose (Bio-Rad Laboratories, Richmond, California) in 0.6 M KC1 buffered to pH 7.4 with 0.02 M Tris- HCI. Wells were filled several times and allowed to diffuse for 7-10 d at 4°C. The plates were then washed with 0.6 M KCl for 5 d, distilled water for 3 d, and 50% alcohol for I d, dried at 37°C and stained with Amido Black. The amount of antigen used was I mg protein/ ml in 0.6 M KCl.

ImmtrnofIuoresccnce microscopy. Tissues were prepared for indirect immunofluorescence microscopy and the microscopy and photography performed as described previously (Jaffe et a!, 1973).

Preparation of antisera for immunopuoresccnce microscopy. The antisera were routinely ab- sorbed with group A, Rh positive human red blood cell stroma, and with platelet poor plasma. For these and subsequent absorptions, the concentration of absorbing protein was I mg/ml.

Sodium dodecyl sulphate (SDS) polyacrylamide gel electrophoresis. SDS polyacrylamide gel electrophoresis ( 5 % gels) was performed as described by Weber & Osborn (1975). Samples for analysis were added to an equal volume of a solution containing 8.5 M urea, 2% SDS, and 14 mM DTT and boiled for 5 min. The gels were stained with Coomassie Brilliant Blue. Densitometric scans of gels were carried out in a Gilford Model 240 spectrophoto- meter equipped with a gel scanning attachment (Gilford Instrument Laboratories, Inc., Oberlin, Ohio) and a Densicord recorder equipped with an integrator (Photovolt Corp., New York).

The molecular weight of myosin was determined by SDS polyacrylamide gel electro- phoresis as described (Weber & Osborn, 1975). Molecular weight markers included : phosphorylase A, mol wt 94 ooo (Sigma Chemical Co.); ovalbumin, mol wt 43 ooo (Phar- macia) ; chymotrypsin, mol wt 25 000 (Calbiochem, San Diego, California) ; and az-macro- globulin subunit, mol wt 185 ooo (kindly provided by Dr Peter Harpel).

RESULTS Endothelial Cell Myosin

A concentrated aliquot of a 0.6 M KCl extract of cultured endothelial cells was chromato-

74 Anne Moore et a/

Volume (ml)

FIG I. Sepharose 4B chromatograph of a 0.6 M KCl extract of cultured human endothclial cells. The myosin-like ATPase activity is localized to the void volunic. 0, Ca2 ATPase; A, EDTA ATPase; m, Mg2 ATPase; - - - -, protein (% transmission at 280 nm).

TABLE I. Purification of myosin from cultured endothelial cells

Sample* Specific ATPase activity?

Ca2+ EDTA Mg2+

Extract supernatant 0.016 0.0028 i0.01

Purified myosin 0.072 0.054 0.014

* Endothelial cells from 33 Petri dishes were extracted with Buffer A yielding XI nig protein. The extract supernatant was divided and 5 . 5 mg was dialysed against Buffer B and assayed for ATPase activity. The remaining 5.5 mg was chromatographed on Sepharose 4B, eluted with Buffer B, and the column eluate assayed for ATPase activity. The purified myosin (600 pg protein) refers to the void volume peak where the ATPase activity was localized.

t pmoles of inorganic phosphorus released/mg protein /min.

graphed on a Sepharose 4B column and thecolumn eluate was monitored for ATPase activity. The calcium and EDTA-stimulated, magnesium-inhibited ATPasc activity was localized to the void volume. The chromatograph of a typical preparation is shown in Fig I. The predominant protein in the void volume peak as analysed by SDS polyacrylamide gel electrophoresis, had a molecular weight of 200 ooo and migrated to the same position on the gel as rabbit uterine myosin heavy chain (Fig 2). The void volume protein also contained a small amount of material of molecular weight 42 000, presumably endothelial cell actin.

Myosin in Endothelial Cells 75

The specific ATPase activities of the endothelial cell myosin before and after column chromatography were compared. Table I shows the ATPase specific activities obtained during a typical purification of endothelial cell myosin. There was a 4+-f0ld increase in calcium ATPase activity and a 20-fold increase in EDTA ATPase activity during the purifica- tion procedure.

Platelet Myosin For comparative purposes, human platelet myosin was partially purified. Platelet extract

was chromatographed on a 4% agarose column (Bio-Gel A-r sm). The myosin-like ATPase activity was localized to the 200-275 ml region of the column (Fig 3). The ATPase specific activity of one platelet myosin preparation is shown in Table 11. The results are similar to those reported by Pollard et a1 (1974). The 200-275 ml fraction, analysed by SDS poly- acrylamide gel electrophoresis, contained a 200 ooo molecular weight protein which migrated

Volume (mi)

FIG 3. Bio-Gel A-ISm chromatograph of platelet extract prepared as in text. The myosin-like ATPase activity is localized to the 200-275 ml fractions. 0, Caz+ ATPase; A, EDTA ATPase, W, MgZ ATPase; - - - -, protein (% transmission at 280 nm).

TABLE 11. Platelet myosin ATPase activity

Sample* Specific ATPase activityt

Caz+ EDTA Mgzt

Platelet myosin 0.83 0.84 <O.OI

* 140 units of platelets yielded 2160 mg of extracted protein. The 1.0-1'9 M ammonium sulphate precipitate represented 17 mg of protein which was chromatographed and yielded 6.6 mg of pro- tein in the zoo-a7j ml eluate.

pmoles inorganic phosphorus released/mg proteinlmin.

76 Anne Moore et al

identically to a rabbit myosin heavy chain marker. The protein in the 200-275 ml region of the column was dialysed against water, lyophilized, and used as an antigen to make antiserum against platelet myosin.

Immunodi@usion Tests The antiserum to platelet myosin formed a line of identity against purified platelet myosin

and crude platelet myosin extract. This antiserum to platelet myosin and an antiserum to human uterine actomyosin also precipitated the identical antigen in the crude platelet myosin preparation (Fig 4). The rabbit antibody to human uterine actomyosin has been previously documented to be specific for actomyosin of the smooth muscle type (Becker & Nachman, 1973), and it is highly probable that this antibody is specific for the myosin moiety. These studies thus demonstrated that the antiserum raised to platelet myosin was specific for platelet myosin.

The antiserum to platelet myosin formed a precipitin line when tested against a crude preparation of endothelial cell myosin, the 0.6 M KCl extract supernatant of endothelial cells (Fig 5 ) . This observation indicated that the endothelial cell myosin antigen was similar or perhaps identical to platelet myosin. Attempts to raise a precipitating antibody to endo- thelial cell myosin have not been successful to date.

None of the antisera formed precipitin lines against normal human platelet poor plasma.

Immunojluorescence Studies The antiserum to platelet myosin intensely stained both endothelial and arteriolar smooth

muscle cells in human tissue sections. This antiserum did not stain cardiac and skeletal muscle. The antiserum to platelet myosin brightly stained cultured endothelial cells in a fine fibrillar pattern (Fig 6). The fibrils appeared to be intracellular and extended along the long axis of the cell. The fibrillar staining disappeared when the antiserum was absorbed with purified platelet myosin and was markedly reduced when the antiserum was absorbed with purified uterine actomyosin. In addition to the fibrillar pattern, there was significant granular perinuclear fluorescence. This perinuclear staining persisted after absorption of the antiserum with platelet myosin. A similar perinuclear pattern has been seen in fibroblasts stained with antiserum to chicken gizzard myosin (Weber & Groeschel-Stewart, 1974). However, endo- thelial cells stained with an antiserum to human uterine actomyosin did not show true perinuclear fluorescence (Becker et 02, 1974). The nature of the perinuclear fluorescence is not known. While it probably represents non-specific staining, there is the possibility that it represents an incomplete ‘pro’-myosin antigen. When the antiserum to endothelial cell myosin was tested with cultured endothelial cells, only faint fibrillar staining of the endo- thelial cells was seen. Normal rabbit serum gave no immunofluorescent staining with any of the tissues mentioned above.

DISCUSSION

Myosin has been isolated from cultured human endothelial cells. The specific activity of the endothelial cell myosin ATPase in the presence of calcium or EDTA was low when compared to the platelet myosin or to other vertebrate non-muscle myosins such as guinea-pig granu-

Myosin in Endothelial Cd l s

FIG 4

FIG 2 FIG 5 FIG 2. SDS polyacrylaiiiidc gels of A, the Scpharosc 4B void volume peak from the endothelial cell preparation and B, purified utcrine actornyosin. The major band in A (arrow) niigratcs in the mole- cular weight range of 200 ooo corrcsponding to the rabbit uterine muscle myosin heavy chain in B. FIG 4. Imniunodiffusion platc dcnionstrating that antiscrum to human platelet myosin (2) and anti- seruni to rabbit utcrinc actoniyosin (3) detcct thc idcntical antigcn in a preparation of crude platelet myosin ( I ) .

FIG 5 . Imniunodiffusion plate demonstrating that the rabbit antiserum to platelet myosin (I) fo rm a prccipitin line with a crude cndothelial cell inyosin extract (2). The stained precipitin line is seen adjacent to wcll I .

(Fncirrg p 76)

Atitre M o m ct n /

FIG 6. Imniunofluorcsccncc study of cultured huinan endothelial cells trcated with rabbit antiscruiii to platelet myosin followed by fluorcsceinatcd goat antiserum to rabbit Ig G. A fibrillar staining pattern was noted throughout the cytoplasm.

Myosin in Endothelial Cells 77 locytes (Stossel & Pollard, 1973), rabbit alveolar macrophages (Hartwig & Stossel, 1975), and rodent fibroblasts (Ostlund et al, 1974). The reason for this comparatively low ATPase specific activity may be related to the lability of the enzyme in the endothelial cell prepara- tion. It is also possible that the comparatively low ATPase activity reflects a relatively 'sluggish' rate of contraction of the endothelial cells in comparison to the brisk activity presumably required of other cells such as macrophages during the process of phagocytosis or of platelets undergoing shape change and aggregation following contact with various aggregating agents. There is strong evidence in skeletal muscles that ATPase activity and speed of muscle shortening are correlated (BArAny, 1967).

The molecular weight of the endothelial cell myosin heavy chain was 200 ooo as deter- mined by its mobility in SDS polyacrylamide gels. Endothelial cell myosin consistently eluted in the void volume of a Sepharose 4B column which has an exclusion limit of twenty million, Fibroblast myosin extracted exactly in the same manner as the endothelial cell myosin eluted immediately after the void volume on a Sepharose 4B column (Ostlund et al, 1974). It is possible that the elution pattern of endothelial cell myosin was due to the presence of myosin aggregates despite the high ionic strength provided by the 0.6 M KCl. The differences in ATPase specific activities and aggregate forming tendencies among the various cytoplasmic myosins suggests that myosins from different sources may be structurally distinct proteins.

Previous immunofluorescence studies from this laboratory showed that endothelial cells, both in tissue sections and in culture, stained with antiserum to uterine smooth muscle actomyosin but not with antiserum to cardiac or skeletal muscle actomyosin (Becker & Nachman, 1973 ; Jaffe et al, 1973). Our present studies provide further evidence that cultured endothelial cells contain a myosin of smooth muscle type. An antiserum specific for platelet myosin detected myosin antigen in an endothelial cell extract (Fig 5) . Further immunologic evidence for the presence of myosin in cultured endothelial cells was provided by immuno- fluorescence studies. Smooth muscle myosin was localized in endothelial cell fibrils which extended along the longitudinal axis of the cultured cell (Fig 6) . This pattern was visualized not only with this antiserum to purified myosin but also has been seen with an antiserum to uterine actomyosin (Becker et al, 1974). The staining pattern is strikingly similar to that reported for fibroblast myosin (Weber & Groeschel-Stewart, 1974).

The immunofluorescence results correlate with the fine structure of the endothelial cell. Bundles of fine filaments have been seen by transmission electron microscopy in cultured human umbilical vein endothelial cells (Jaffe et al, 1973). Cloned guinea-pig portal vein endothelial cells, which contain a myosin-like protein on SDS gel electrophoresis, also contain filament bundles which are located on the attachment side of the cells (Blose, 1976). Thick and thin filament bundles have been identified by electron microscopy in vertebrate coronary artery endothelium (Yohro & Burnstock, 1973). Filament bundles have been seen in rat splenic sinusoid endothelium (De Bruyn & Cho, 1974). In addition, rat arterial endothelial cells appear to contain filaments which can be decorated by heavy meromyosin suggesting that actin filaments exist in the endothelial cell (Rostgaard et al, 1972).

The amount of myosin in cultured endothelial cells based on densitometric scanning of the protein fraction with maximum ATPase activity was approximately O . S - I ~ ~ of the total cell protein. In comparison, human platelets contain about 1.5% myosin (Pollard et al, I974),

78 Anne Moore et a1

rabbit alveolar macrophages contain 0.7-1.5% myosin (Hartwig & Stossel, 1975), and cultured rodent fibroblasts contain about 0.5-3% myosin (Ostlund et al, 1974).

The localization of myosin in non-muscle cells has been studied by, a number of invest- igators. Although the platelet was one of the earliest non-muscle cells in which contractile proteins were identified (Bettex-Galland & Liischer, 1959), the question of the intracellular location of platelet myosin has not been resolved (Pollard & Weihing, 1974). Studies of the localization of myosin in cultured cells using immunologic techniques have yielded differing results. Willingham et al (1974) studied the distribution of myosin in mouse L929 cells using specific antiserum to L929 cell myosin. Their results suggested that some myosin was localized to the cell surface. Weber & Groeschel-Stewart (1974) studied myosin in cultured fibroblasts with indirect immunofluorescence using an antiserum to chicken gizzard myosin. They described a complex network of fibrils which spanned the interior of the cell and which appeared to lie toward the adhesive side of the cell. These authors concluded that the myosin-containing filaments were closely related to or even identical to actin-containing filaments which have also been' visualized in cultured fibroblasts by indirect immuno- fluorescence (Lazarides & Weber, 1974). Our results are similar to those reported for the fibroblast and demonstrate an intracellular filamentous localization for myosin in cultured endothelial cells.

Myosin is found in a variety of vertebrate and invertebrate non-muscle cells and may be important in activities such as primitive locomotion, cell division, and phagocytosis (Pollard & Weihing, 1974). The discovery of a smooth muscle myosin in human endothelial cells is not surprising in view of the previously described contractile behaviour of these cells (Majno et a!, 1969). It has been hypothesized that endothelial cell contraction may be im- portant in such diverse processes as the regulation of vascular permeability in response to inflammatory stimuli, the contact of platelet and subendothelial collagen with subsequent initiation of haemostasis (Becker & Nachman, 1g73), and the deposition of immune com- plexes along vascular basement membranes (Cochrane & Koffler, 1973).

The antigenic similarity between the platelet and the endothelial cell contractile systems is intriguing and is further support for the concept that the endothelial cell and the platelet are intimately related. Previous studies have suggested that platelets are necessary for the preservation of normal vascular integrity both in viuo (Gimbrone et al, 1969; Roy & Djerassi, 1972), and in uitro (Saba & Mason, 1975). In addition, ultrastructural studies of experimental thrombocytopenic purpura in guinea-pigs have shown widened intercellular junctional spaces in the endothelial lining of large capillaries and venules (Gore et al, 1970). A role for the endothelial cell in the support of normal platelet function has been demon- strated by the recent report of the synthesis of von Willebrand factor by cultured human endothelial cells (Jaffe et a2, 1974). It is likely that more precise characterization of the con- tractile protein systems in endothelial cells and platelets will clarif) the interaction of these cells in physiologic as well as certain pathologic processes.

ACKNOWLEDGMENTS

This investigation was supported by National Institutes of Health Grants HL-14810 and HL-1803 and grants from The Amerio Foundation, The Arnold R. Krakower Foundation, The National Hemophilia Foundation and The Cross Foundation.

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