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Previous Volume 340:994-1004 April 1, 1999 Number 13 Next

Abstract PDF Editorial by Rand, J. H. Add to Personal Archive Add to Citation Manager Notify a Friend E-mail When Cited PubMed Citation

ABSTRACT

Background Acute promyelocytic leukemia (APL) is associated with a hemorrhagic disorder of unknown cause that responds to treatment with all-trans-retinoic acid.

Methods We studied a newly described receptor for fibrinolytic proteins, annexin II, in cells from patients with APL or other leukemias. We examined initial rates of in vitro generation of plasmin by tissue plasminogen activator (t-PA) in the presence of APL cells that did or did not have the characteristic translocation of APL, t(15;17). We also determined the effect of all-trans-retinoic acid on the expression of annexin II and the generation of cell-surface plasmin.

Results The expression of annexin II, as detected by a fluorescein-tagged antibody, was greater on leukemic cells from patients with APL than on other types of leukemic cells (mean fluorescence intensity, 6.9 and 2.9, respectively; P<0.01). The t(15;17)-positive APL cells stimulated the generation of cell-surface, t-PA–dependent plasmin twice as efficiently as the t(15;17)-negative cells. This increase in plasmin was blocked by an anti–annexin II antibody and was induced by transfection of t(15;17)-negative cells with annexin II complementary DNA. The t(15;17)-positive APL cells contained abundant messenger RNA for annexin II, which disappeared through a transcriptional mechanism after treatment with all-trans-retinoic acid.

Conclusions Abnormally high levels of expression of annexin II on APL cells increase the production of plasmin, a fibrinolytic protein. Overexpression of annexin II may be a mechanism for the hemorrhagic complications of APL.

Another feature of APL is a hemorrhagic diathesis, which is thought to result from disseminated intravascular coagulation, abnormal fibrinolysis, or both.¹ Evidence of enhanced thrombin activation supports the mechanism of disseminated intravascular coagulation.^{12,13,14,15,16} However, plasma levels of the anticoagulant proteins antithrombin III and protein C are usually normal^17,18,19 and platelet survival is normal²⁰ in patients with APL, unlike the findings in patients with disseminated intravascular coagulation.

Plasminogen and its activators, tissue plasminogen activator (t-PA) and urokinase plasminogen activator (u-PA), generate plasmin, a proteolytic enzyme that cleaves fibrinogen and fibrin, thereby dissolving clots.²¹ Overproduction of this fibrinolytic enzyme can cause abnormal bleeding. The hemorrhagic complications of APL may be due to increased fibrinolysis. Evidence in support of this possibility includes low plasma levels of plasminogen, ₂-plasmin inhibitor (the primary plasmin inhibitor), plasminogen-activator inhibitor 1 (an inhibitor of both t-PA and u-PA), and other abnormalities in patients with APL that are consistent with excessive fibrinolysis.^{12,13,18,22,23,24,25,26,27,28,29,30,31,32}

Annexin II is a calcium-regulated, phospholipid-binding protein on endothelial cells, macrophages, and some tumor cells.³³ It is a cell-surface receptor for both plasminogen (the inactive precursor of plasmin), and its activator, t-PA.³⁴ Soluble annexin II acts as a t-PA cofactor, increasing the efficiency of plasmin formation by a factor of 60.³⁵

We examined the expression of annexin II on leukemic cells from 14 patients with APL or other leukemias and determined the initial rate of plasmin generation in myeloid leukemic cells that were positive or negative for the t(15;17) translocation. We correlated plasmin activity with the expression of both annexin II protein and messenger RNA (mRNA). All-trans-retinoic acid reversed the excessive annexin II–mediated fibrinolytic activity of leukemic promyelocytes by blocking transcription of the annexin II gene in translocation-positive cells. This effect of the drug may explain the reversal of the bleeding tendency in APL within the first days of treatment with all-trans-retinoic acid.

Methods

Isolation of Leukemic Cells

Surplus peripheral-blood leukocytes or bone marrow samples from 14 patients with leukemia were treated with heparin, coded to maintain the patients' anonymity, and centrifuged with Ficoll–Hypaque (Sigma, St. Louis) at 800xg for 30 minutes. Cells were collected at the interface between the Ficoll–Hypaque and HEPES-buffered saline, washed in RPMI 1640 medium, and resuspended in growth medium.

Cell Culture

NB4 cells, a stable, translocation-positive cell line, were cultured as described elsewhere³⁶; the cells were provided by Dr. M. Lanotte (Hôpital St. Louis, Paris). Human umbilical-vein endothelial cells were propagated as described elsewhere.³⁷ Cells from a patient with acute myeloblastic leukemia (AML) characterized by poorly differentiated myeloblasts (AML-M1) were provided by Dr. S. Rafii (Weill Medical College of Cornell University, New York). APL-1 cells, cloned from a patient with APL, were shown to be t(15;17)-negative by fluorescence in situ hybridization (performed by Dr. M.J. Macera, Long Island College Hospital, Brooklyn, N.Y.) and by a reverse-transcriptase–polymerase-chain-reaction assay (performed by Dr. E. Dmitrovsky, Memorial Sloan-Kettering Cancer Center, New York).³⁸ HL-60 cells derived from a patient with AML were provided by Dr. P. Tempst (Memorial Sloan-Kettering Cancer Center).^39,40 All leukemic cells were propagated in RPMI 1640 medium containing 10 percent fetal-calf serum, 2 mM glutamine, penicillin (100 U per milliliter), streptomycin (100 µg per milliliter), and amphotericin B (0.25 µg per milliliter).

Flow Cytometry

Washed cells from 13 patients were incubated with rabbit preimmune or anti–annexin II IgG³⁵ (100 µg per milliliter) for 15 minutes at 4°C, washed three times, incubated with fluorescein isothiocyanate–conjugated goat antirabbit IgG (20 µg per milliliter) for 30 minutes at 4°C, fixed in 2 percent paraformaldehyde for 2 minutes at 21°C, and analyzed on an Epics flow cytometer (Coulter, Miami). With the use of lysates from human umbilical-vein endothelial cells and NB4 cells, the rabbit anti–annexin II IgG and a mouse monoclonal IgG antibody specific for annexin II reacted with the same single band on Western blotting.

Indirect Immunofluorescence Microscopy

Cells were centrifuged onto cytospin slides (134xg) for 6 minutes at 21°C, air dried, and fixed with 3.7 percent formaldehyde for 20 minutes at 21°C. The slides were washed, blocked with 0.1 percent bovine serum albumin and 1 percent normal goat serum in Dulbecco's phosphate-buffered saline for 20 minutes at 21°C, washed again, incubated with polyclonal anti–annexin II IgG or control rabbit preimmune IgG (12 to 24 µg per milliliter) for 1 hour at 21°C, and incubated with fluorescein isothiocyanate–conjugated goat antirabbit IgG (8 µg per milliliter) for 1 hour at 21°C. The slides were then washed five times and counterstained with Evans blue or propidium iodide.

Plasminogen Activation Assay

Cells were preincubated with 100 nM lysine–plasminogen or 200 nM glutamic acid–plasminogen (Immuno, Vienna, Austria) for one hour at 21°C. Then, 10 nM t-PA (Genentech, South San Francisco, Calif.) and the plasmin substrate d-valine-leucine-lysine-7-amino-4-trifluoromethyl coumarin (AFC-81, Enzyme Systems Products, Dublin, Calif.) were mixed and added in the presence or absence of the following inhibitors: amiloride (Sigma), anti-urokinase IgG (no. 3940A, American Diagnostica, Greenwich, Conn.), anti–t-PA IgG (no. 364B, American Diagnostica), anti–annexin II IgG (Oncogene Research Products, Cambridge, Mass.), and anti–annexin I IgG (Zymed, San Francisco). Substrate cleavage was measured in duplicate or triplicate at two-minute intervals (excitation, 400 nm; emission, 505 nm) with 2-nm slit widths in a fluorescence spectrophotometer (model 650-10S, Perkin-Elmer, Norwalk, Conn.) as described elsewhere.³⁵ Anti–annexin I IgG and anti–annexin II IgG were pretreated with carboxypeptidase B–sepharose as described elsewhere.⁴¹

Ribonuclease Protection Assay

Bases 51 through 350, encoding the unique "tail" region of annexin II,⁴² were amplified by the polymerase chain reaction from human annexin II complementary DNA (cDNA)³⁴ with the use of primers 5'AAAGGATCCTGTCTACTGTTCACG3' and 5'AAAGAATTCCCAAAATCACCGTCT3', ligated into pBluescript KS(+) at EcoRI and BamHI restriction sites,⁴³ and propagated in transformed Escherichia coli selected on the basis of its resistance to ampicillin. Plasmids were isolated with the Maxi-Prep kit (Qiagen, Chatsworth, Calif.), linearized with EcoRI, and purified. Radiolabeled probes for annexin II and glyceraldehyde phosphate dehydrogenase (Amersham, Arlington Heights, Ill.) were transcribed with an RNA-transcription kit (Stratagene, La Jolla, Calif.) and the use of [³²P]uridine triphosphate and either T3 or T7 RNA polymerases to yield 377-base and 139-base antisense riboprobes with 300-base and 100-base targets, respectively. Hybridizations were carried out with the Direct Protect kit (Ambion, Austin, Tex.). Double-stranded protected RNA fragments were visualized by autoradiography of dried gels (7 M urea, 6 percent polyacrylamide, and TRIS, borate, and EDTA buffer) and analyzed by densitometry or quantitated with a Phosphorimager (Molecular Dynamics, Sunnyvale, Calif.).

Nuclear Run-On Assays

The pBluescript vector containing 300 bases of the annexin II tail sequence and a pBR322 plasmid containing 4.5 kb of a 28S ribosomal RNA sequence (provided by Dr. I. Gonzales, Hahnemann Hospital, Philadelphia) were used to screen radiolabeled transcription products in nuclear run-on assays.⁴⁴ Plasmids were linearized and applied to nitrocellulose membranes with the use of a vacuum slot-blot apparatus (Hoefer Instruments, San Francisco). Radiolabeled transcripts were purified as described elsewhere⁴⁴ and hybridized at 65°C for 36 hours.

Transfection of APL-1 Cells

APL-1 cells (2x10⁶ per milliliter) were incubated with 20 µg of Lipofectin per milliliter (Life Technologies, Gaithersburg, Md.) and an annexin II–expression vector, pCMV5-Annexin II (2 µg per milliliter for 24 hours); supplemented with minimal essential medium; and assayed for plasmin at 48 hours.³⁴ The efficiency of transfection was assessed on the basis of the expression of ß-galactosidase activity after the introduction of pSV-ß-galactosidase (Promega). The empty pCMV5 vector served as the control.

Statistical Analysis

Data were analyzed with Student's two-tailed t-test.

Results

Patients

All six patients with APL had evidence of increased fibrinolysis (low plasma levels of fibrinogen, high plasma levels of fibrin split products, or high plasma levels of d-dimer, alone or in combination) (Table 1). In the four patients with APL who also had severe thrombocytopenia (platelet count, <20,000 per cubic millimeter), there was overt bleeding, including a life-threatening pulmonary hemorrhage in one case. In Patient 2, who had minimal bleeding at presentation, fibrinogen levels decreased during cytotoxic chemotherapy. This patient required a continuous infusion of aminocaproic acid and heparin. Severe hypofibrinogenemia and an elevated plasma d-dimer level developed in Patient 5, who had APL that was resistant to treatment with all-trans-retinoic acid, and he died from multiorgan failure. (The cells from this patient were tested one week after treatment with all-trans-retinoic acid had been discontinued.) Patient 6 presented with relapsed APL and received arsenic trioxide. A pulmonary hemorrhage and a coagulation disturbance improved during the first week of therapy; similar results have been reported previously.⁴⁵

View this table: [in this window] [in a new window]   Table 1. Clinical Findings and Annexin II Expression in 13 Patients with Leukemia.

 
Annexin II Expression in APL Cells

Immunofluorescence microscopy was used to detect annexin II in promyelocytes from patients with APL (Figure 1A, Figure 1B, Figure 1C, Figure 1D, and Figure 1E). The results were strongly positive with t(15;17)-positive leukemic blasts (Figure 1A and Figure 1C), whereas the results were only slightly positive with the t(15;17)-negative cell line, APL-1 (Figure 1E). Staining of (t15;17)-positive APL cells with IgG from the serum of an unimmunized rabbit was negative (Figure 1B). Leukemic blasts from a patient with AML characterized by minimal myeloid differentiation did not react with the anti–annexin II antibody (Figure 1D).

View larger version (559K): [in this window] [in a new window]   Figure 1. Immunofluorescence Staining of APL Cells with Anti–Annexin II Antibodies. Cells from two patients with t(15;17)-positive APL (Panels A and C) and one patient with AML characterized by minimal myeloid differentiation (Panel D) and cells from the t(15;17)-negative APL cell line (APL-1) (Panel E) were stained with anti–annexin II (Panel A, x600; Panels C, D, and E, x1000). The t(15;17)-positive APL cells shown in Panel A were also stained with preimmune IgG (Panel B, x600). Cells were counterstained with propidium iodide (Panels A, B, D, and E) or Evans blue (Panel C).

 
Flow-cytometric studies showed that t(15;17)-positive APL cells from three patients with recent diagnoses and three with relapses had mean fluorescence intensities with anti–annexin II antibodies that were 3.8 to 10.9 times the intensity observed with the control IgG (Table 1). Leukocytes from six of the seven patients with other forms of leukemia expressed lower levels of annexin II. The relative mean fluorescence intensity for APL cells was 6.9, whereas for AML and acute lymphocytic leukemia cells it was 2.9 (P<0.01). One patient with relapsed AML (Patient 13) had a relatively high level of annexin II (mean fluorescence intensity, 5.9). She also had elevated plasma levels of d-dimer, a prolonged prothrombin time, severe thrombocytopenia, and excessive bleeding.

The expression of annexin II on t(15;17)-positive APL cell lines was further evaluated by Western blotting of eluates from cell surfaces.⁴⁶ With the use of an annexin II–specific monoclonal IgG antibody, annexin II was detected in eluates from umbilical-vein endothelial cells and t(15;17)-positive NB4 cells but not in eluates from t(15;17)-negative cell lines (HL-60, AML-M1, and APL-1) (data not shown).

Generation of Plasmin in APL Cells

Using a fluorogenic assay, we assessed the ability of t(15;17)-positive NB4 cells to activate plasminogen (Table 2). By itself, t-PA was a weak plasminogen activator, but with t-PA in the presence of NB4 cells, the rate of plasmin generation was increased by a factor of 28. Approximately 45 percent of this increment occurred in the absence of t-PA, whereas about 55 percent was dependent on exogenous t-PA (P<0.001). Without added t-PA, plasminogen activation was inhibited by the u-PA–specific antagonist amiloride and by anti–u-PA antibodies but not by anti–t-PA antibodies. These results suggest that endogenous production of plasmin by NB4 cells is largely due to u-PA and that NB4 cells enhance plasminogen activation by mechanisms that depend on t-PA and by mechanisms that are independent of t-PA.

View this table: [in this window] [in a new window]   Table 2. Plasminogen Activation in the Presence of Leukemic Cells.

 
NB4 cells stimulated t-PA–dependent activation of plasminogen more effectively than equivalent numbers of t(15;17)-negative cells (Table 2). For HL-60, AML-M1, and APL-1 cells, the rates of activation were 58.8, 48.2, and 45.2 percent of the value obtained with NB4 cells (P<0.001), respectively, suggesting that cells with the t(15;17) translocation support plasmin generation much more efficiently than cells without the translocation. Furthermore, in the presence of aminocaproic acid, a lysine analogue that inhibits the binding of plasminogen to annexin II,³⁷ plasmin production by NB4 cells was reduced to 29.2 percent (P<0.001) of the rate in its absence, whereas the formation of plasmin in the soluble phase was not affected. Moreover, a monoclonal anti–annexin II antibody reduced plasmin generation by NB4 cells to 65 percent (P<0.02) of that observed when NB4 cells were incubated with an equivalent concentration of anti–annexin I IgG antibody. These results constitute evidence that annexin II on the surface of leukemic promyelocytes has a key role in the production of plasmin.

For further analysis of the role of annexin II in the activation of plasminogen by t-PA, t(15;17)-negative cells (APL-1), which lack cell-surface annexin II, were transfected with a plasmid containing either the full-length annexin II cDNA or an empty vector.³⁴ APL-1 transfectants stimulated plasmin production 2.7 times as effectively as nontransfected cells (P<0.001) and nearly twice as effectively as cells transfected with the empty vector (P<0.01). These data suggest that the expression of annexin II is directly correlated with the capacity to generate plasmin.

Annexin II mRNA levels were determined with a ribonuclease protection assay. Steady-state mRNA levels in two t(15;17)-negative cell lines, AML-M1 and APL-1, were 11 and 10 percent, respectively, of levels in t(15;17)-positive APL cells (NB4 cells) (Figure 2). HL-60 cells expressed nearly equivalent levels of annexin II mRNA (74 percent), but the protein was not detected on the cell surface in experiments involving ethylene glycol-bis(ß-aminoethyl ether)-N,N,N',N'-tetraacetic acid elution.

View larger version (73K): [in this window] [in a new window]   Figure 2. Steady-State Levels of Annexin II mRNA. Guanidine thiocyanate lysates from five cell types were hybridized with annexin II and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) riboprobes to yield protected fragments of 300 and 100 bases, respectively. Probes before and after treatment with ribonuclease, as well as protected fragments from each sample, are shown. The numbers at the bottom of the figure indicate the quantities of the annexin II mRNA fragment, normalized to the GAPDH fragment, expressed as the value relative to that in NB4 cells. HUVEC denotes human umbilical-vein endothelial cells. These cells are included as a positive control for the expression of annexin II mRNA.

 
Effect of All-trans-Retinoic Acid on the Synthesis of Annexin II by NB4 Cells

After five and seven days of treatment of NB4 cells with all-trans-retinoic acid, the rate of plasmin production fell to nearly that observed in t(15;17)-negative cells (61.1 and 67.5 percent, respectively, of the rate in untreated control cells; P<0.001). Similarly, cellular expression of annexin II in cells treated with all-trans-retinoic acid was lower than that in mock-treated controls at 72 hours and was completely absent at 120 hours (Figure 3A).

View larger version (152K): [in this window] [in a new window]   Figure 3. Expression of Annexin II Protein (Panel A) and Annexin II Messenger RNA (mRNA) (Panel B) in NB4 Cells. To study the effect of all-trans-retinoic acid on the expression of annexin II protein, we treated NB4 cells with 0.1 percent ethanol (control cells) or 1 µM all-trans-retinoic acid and then incubated the washed cells in calcium-free Dulbecco's phosphate-buffered saline and 10 mM ethylene glycol-bis(ß-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) for 30 minutes at 4°C. Panel A shows Western blots of EGTA extracts of NB4 cells with the use of monoclonal anti–annexin II IgG after treatment with ethanol (control, C) or all-trans-retinoic acid (R) for the indicated periods. The molecular-size marker (mass ratio, Mr) and native annexin II (25 µg) are shown in the first and last lanes, respectively. The membrane was probed again for glyceraldehyde-3-phosphate dehydrogenase (GAPDH) with the use of a monoclonal antibody from Biodesign International (Kennebunk, Me.) and detected with the use of the Enhanced ChemiLuminescence kit (Amersham). Panel B shows the results of the ribonuclease protection assay for annexin II mRNA. Lysates of NB4 cells were treated for the indicated periods with 0.1 percent ethanol (C) or 1 µM all-trans-retinoic acid (R).

 
The effect of all-trans-retinoic acid on annexin II mRNA levels in NB4 cells was evaluated by a ribonuclease protection assay (Figure 3B). Annexin II mRNA levels were reduced within 24 hours after treatment with all-trans-retinoic acid and reached a nadir after 72 hours. The inhibitory effect of all-trans-retinoic acid was evident over a range of concentrations (50 percent inhibitory concentration, <10 nM). For concentrations of 100 nM or more, the rate of inhibition was greater than 85 to 90 percent. Moreover, the expression of annexin II mRNA was diminished after treatment for 48 hours with a stereoisomer of all-trans-retinoic acid, 13-cis-retinoic acid (36 percent of the control value), but not on exposure to two non–retinoid-differentiating agents, phorbol myristate acetate⁴⁷ and vitamin D₃ (87 and 97 percent of the control value, respectively). These data indicate that the expression of annexin II in NB4 cells exhibits a high degree of retinoid-specific sensitivity.

Using nuclear run-on analyses, we assessed the effect of all-trans-retinoic acid on the rate of annexin II gene transcription. In two experiments, the average rate of annexin II transcription was reduced to 70 and 80 percent of the value in vehicle-treated controls after 12 and 24 hours of treatment with all-trans-retinoic acid, respectively. In three experiments, the mean (±SE) rate was reduced to 39±8 percent of the value in controls after 48 hours (P<0.001). Similarly, NB4 cells treated with all-trans-retinoic acid were exposed to dactinomycin (2 µg per milliliter), and steady-state annexin II mRNA levels were estimated with the ribonuclease protection assay. In four separate experiments, no increase in mRNA degradation was observed after three or six hours of treatment with dactinomycin. These data indicate that the inhibition of annexin II expression by all-trans-retinoic acid is transcriptionally mediated and not the result of accelerated degradation of annexin II mRNA.

Discussion

These studies showed that APL cells with the t(15;17) translocation expressed abnormally high levels of cell-surface annexin II. The cells supported rapid rates of plasmin production by t-PA, an effect that was inhibited by anti–annexin II antibodies. Moreover, t(15;17)-negative APL-1 cells that were transfected with annexin II cDNA also exhibited increased t-PA–dependent production of plasmin. These data suggest that overexpression of annexin II increases plasmin production by t(15;17)-positive APL cells, thereby contributing to the hemorrhagic diathesis of APL. Although the presence of the t(15;17) translocation appears to be correlated with overexpression of annexin II, further studies are warranted to verify this finding.

In patients with APL, bleeding usually begins to resolve within five to seven days after the start of treatment with all-trans-retinoic acid, and plasma levels of fibrinogen and ₂-plasmin inhibitor return to normal.¹³ In our series, plasma fibrinogen levels returned to normal within the first week of therapy in two of the three patients with APL who received all-trans-retinoic acid. In vitro treatment of t(15;17)-positive APL cells with all-trans-retinoic acid significantly reduced both the cellular expression of annexin II and plasmin generation over a similar period. Since the half-life of annexin II is approximately 15 hours,⁴⁸ the observed rate of protein disappearance represents between 5 and 8 annexin II half-lives. This time course is consistent with the time that it took for annexin II mRNA to be reduced by 50 percent in the presence of all-trans-retinoic acid in vitro (12 to 18 hours). Because all-trans-retinoic acid did not stimulate the degradation of annexin II mRNA but did impair its production, we conclude that the inhibitory effect of all-trans-retinoic acid on the expression of annexin II is transcriptionally mediated.

Annexin II is thought to have a thromboregulatory role by enhancing the t-PA–dependent formation of plasmin on the endothelial cell surface.³³ Dysregulated expression of annexin II on the surface of circulating APL cells may lead to uncontrolled production of plasmin, thereby shifting the hemostatic balance toward overt bleeding (Figure 4A and Figure 4B). Because plasmin formed on cell surfaces appears to be protected from its primary inhibitor, ₂-plasmin inhibitor,⁴⁹ the total fibrinolytic effect of overexpression of annexin II may be clinically significant. Furthermore, because ₂-plasmin inhibitor may become depleted in patients with APL,^{12,13,18,22,24,25} circulating plasmin may go unchecked, further increasing the potential for hemorrhage.

View larger version (152K): [in this window] [in a new window]   Figure 4. Proposed Mechanism of Hemorrhage in APL. Plasmin is formed on assembly of plasminogen and tissue plasminogen activator (t-PA) on cell-surface–associated annexin II (Panel A).33 At the cell surface, plasmin is protected from its primary inhibitor, 2-plasmin inhibitor (2-PI),49 which is produced in the liver. Once released, plasmin rapidly forms an irreversible, inactive complex with 2-PI.21 Plasmin is generated on the surface of endothelial cells and, to a lesser extent, on other cells. In leukemias other than APL, released plasmin is neutralized by 2-PI, and the plasmin–2-PI complexes are cleared in the liver. In APL, plasmin is generated at an abnormally high rate because of the overexpression of annexin II on the leukemic cells (Panel B). As a result, 2-PI is consumed, and active plasmin accumulates in the plasma. The unopposed fibrinolytic activity of plasmin causes a hemorrhagic disorder.

 
Our in vitro studies of NB4 cells indicate that u-PA, in addition to t-PA, may play a part in the generation of plasmin by APL cells. Urokinase has been reported to be produced by these cells.^26,30,32 In the absence of t-PA, approximately two thirds of the base-line plasmin generation by NB4 cells was inhibited by anti–u-PA antibody and by amiloride (a u-PA–specific inhibitor). The presence of u-PA would explain why approximately 30 percent of the plasmin production was insensitive to the plasminogen-binding inhibitor aminocaproic acid (Table 2).

In addition to annexin II, other annexins may regulate the formation of clots. Annexin VIII is expressed in APL cells and is down-regulated by all-trans-retinoic acid,^50,51 but it is not known whether this intracellular protein affects the hemostatic balance. Annexin V is expressed on the surface of placental villi cells.⁵² In the antiphospholipid antibody syndrome, annexin V appears to be displaced by IgG antiphospholipid antibodies. This process may expose membrane phospholipid, resulting in vascular thrombosis in the placenta and fetal wastage. Thus, the annexins may represent a unique group of proteins that regulate hemostasis.

The leukemic promyelocytes from all six patients with APL in our study expressed high levels of annexin II, and all these patients had evidence of accelerated fibrinolysis. Several small studies have suggested that treatment with antifibrinolytic drugs, such as aminocaproic acid and tranexamic acid, may reduce complications due to bleeding in patients with APL, further supporting the role of the fibrinolytic system in the coagulation disturbance.^24,53,54,55 However, several reports of the development of thromboses in patients with APL during treatment with all-trans-retinoic acid suggest that it should be used cautiously.^56,57,58 Our study included two patients with AML of monocytic lineage, one of whom had severe bleeding complications and a high level of expression of annexin II on leukemic monocytes. We have observed annexin II on the surface of monocyte-derived macrophages,⁵⁹ suggesting that the expression of this protein may not depend solely on the t(15;17) translocation. Nevertheless, the level of annexin II expression by leukemic cells may be a useful factor in deciding whether or not to use antifibrinolytic therapy. Studies currently under way are examining the correlation between the level of annexin II expression and the degree of bleeding and fibrinolytic activity in patients with leukemia.

In summary, a high level of expression of annexin II appears to be a marker of APL and may contribute to bleeding disorders in patients with APL by activating the fibrinolytic system. Treatment with all-trans-retinoic acid down-regulates the production of mRNA for annexin II, which may explain the rapid resolution of coagulopathy in patients receiving retinoid therapy.

Supported by grants from the National Institutes of Health (HL 42493, HL 46403, HL 58981, and HL 03558) and the Robert Steel Foundation for Pediatric Cancer Research.

We are indebted to Drs. R. Warrell, P. Steinherz, A. Aledo, and J. Garvin for assistance in obtaining samples from patients.

Source Information

From the Division of Hematology–Oncology, Departments of Pediatrics (J.S.M., A.T.J., E.A.L., K.A.H.) and Medicine (K.A.H.), Weill Medical College of Cornell University, New York; the Division of Hematology–Oncology, Department of Medicine, Instituto Nacional de la Nutricion Salvador Zubirán, Mexico City, Mexico (G.M.C.); and the Division of Cardiology, Department of Medicine, Mount Sinai Medical Center, New York (M.A.M.).

Address reprint requests to Dr. Menell at Columbia University, College of Physicians and Surgeons, 180 Ft. Washington Ave., HP5, New York, NY 10032, or at menellj{at}sjhmc.org.

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