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* Department of Medicine and Institutes of
Pathology and
Immunology, University of Münster, Münster, Germany
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Abstract |
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Introduction |
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For platelets, it has been shown that a close interaction with and adhesion to neutrophilic granulocytes and monocytes take place depending on the level of cell activation (8, 9). Adherence between monocytes/macrophages and platelets occur in the vessel wall and plaque, but it is also demonstrable in the blood stream where it has been called platelet satellitism (10). This phenomenon has been attributed to thrombotic disorders such as stroke. Also, consequences for leukocyte apoptosis after the interaction with platelets have been studied. In vitro, neutrophils are rescued from apoptosis by platelets by a process that is independent of P-selectin or glycoprotein IIb/IIIa (11). Similarly, mediators stored in platelet granules, such as platelet factor 4, have been shown to support monocyte survival and to induce monocyte differentiation (12). In the present study, the effects of freshly isolated platelets on monocyte senescence were studied in an in vitro system. We could show that spontaneous and CD95-mediated monocyte apoptosis was markedly suppressed by platelets due to rapid phagocytosis of thrombocytes. This phenomenon might have further clinical implications not only in the setting of atherogenesis but also in various other inflammatory disorders.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PE-conjugated mAb against activated caspase-3, anti-caspase-8 polyclonal Ab, and anti-caspase-9 mAb (clone 2-22) was purchased from BD PharMingen (San Diego, CA). Anti-CD95/FAS mAb (clone CH-11, 500 ng/ml) and FITC-labeled anti-CD62P mAb (20–40 µg/ml) were obtained from Coulter Immunotech (Krefeld, Germany). Anti-heat shock protein (hsp-70)3 mAb was purchased from StressGen (Victoria, Canada), reverse transcriptase from Stratagene (Heidelberg, Germany), Taq DNA polymerase from Life Technologies (Karlsruhe, Germany), dNTPs from New England Biolabs (Beverly, MA), pd(N)6 from Boehringer Mannheim (Mannheim, Germany), and agarose from AGS (Heidelberg, Germany). All other reagents including EDTA/trypsin solution were purchased from Sigma-Aldrich (St. Louis, Missouri), unless otherwise indicated.
Monocytes and cell culture
Human monocytes were isolated from leukocyte buffy coats or healthy volunteers. Mononuclear cells were obtained by Ficoll-Hypaque density gradient centrifugation (400 x g, 20 min), washed, and further purified by centrifugation on a hypotonic Percoll density gradient (57% in PBS; 400 x g, 30 min). Two interphases were found in which the upper phase contained the enriched monocytes. Cells were collected, washed three times in cold PBS, and seeded out in 24-well culture plates (Greiner, Nürtingen, Germany) in RPMI 1640 culture medium containing 2 mM L-glutamine, 50 µg/ml penicillin/streptomycin, 5 mM HEPES, and 10 µM 2-ME at 37°C in a 5% CO2/95% air atmosphere. Monocytes were further purified by the adherence to the culture plates, which finally gave a purity of >85%. This was assessed by flow cytometry on a FACScan flow cytometer (BD Biosciences, Mountain View, CA) defined by forward and side light scatter properties, as well as by detection of the CD14 surface molecule and staining for nonspecific esterase. Monocytes (0.5 x 106/ml) were incubated in culture medium with 0.2–5% FCS (endotoxin content, < 0.01 ng/ml) or were rendered apoptotic with a CD95 mAb (0.5 µg/ml) applied over the 60-h culture time.
Preparation of platelets, determination of surface receptors, and coculture with monocytes
Platelets were prepared according to standard methods from 20 ml blood from healthy donors anticoagulated with 0.01 M sodium citrate. Centrifugation was performed at 200 x g for 10 min to get platelet-rich plasma. Platelets were washed first in an apyrase (0.5 U/ml) containing EDTA (5 mM) buffer, then twice in 5 ml HEPES/Tyrode buffer, each with centrifugation at 800 x g for 10 min. Finally, platelets were resuspended in RPMI 1640 containing 0.2% FCS and immediately transferred to monocyte cultures. Platelets (5 x 106–20 x 106/ml) were added to monocytes after 20 h for 4 h at a monocyte:platelet ratio of 1:10–1:40. Platelets kept in culture over this short time period exerted normal shape and function as determined morphologically and by surface receptor analysis. A ratio of 1:10 was applied in all experiments if not otherwise indicated. After vigorously washing monocyte/platelet cocultures to remove unbound platelets, monocytes were further incubated for 36 h to allow progression or inhibition of monocytes. For activation, thrombocytes were prepared in a hypotonic HEPES buffer containing only 0.021 mM NaCl (13). CD62P expression of platelets as an activation marker was determined after fixation in 1% paraformaldehyde for 60 min by flow cytometry using a specific mAb.
Platelet culture supernatants were harvested after a 48-h culture of thrombocytes; platelet granule contents were obtained by ultrasonification.
Detection of apoptosis
Quantification of apoptosis by flow cytometry. Monocyte apoptosis was determined by propidium iodide (PI) staining of nuclei using flow cytometry. This procedure is based on the principle that after DNA fragmentation permeabilized cells exhibit a reduced chromatin stainability and accessibility to fluorochromes (14). Monocytes were washed in PBS, fixed with 4% paraformaldehyde, and permeabilized with 0.1% saponin. For staining, PI (5 µg/ml) was applied for 15 min before cells were washed again in PBS containing 0.01% NaN3. Cells were analyzed on a FACScan flow cytometer (BD Biosciences) for a total of 10,000 events.
TUNEL labeling. DNA strand breaks were identified by labeling free 3'-OH termini with modified nucleotides in an enzymatic reaction using the in situ cell death detection kit, Fluorescein (Boehringer Mannheim) according to the manufacturer’s instructions. In brief, monocytes were fixed in 4% paraformaldehyde at room temperature for 15 min. Subsequently, cell membranes were permeabilized with 0.1% Triton X-100 for 3 min on ice. Then DNA was labeled with FITC-conjugated dUTP in the presence of TdT for 1 h at 37°C. Cells were washed extensively in PBS with 1% BSA and analyzed by flow cytometry or immunofluorescence microscopy. Cells labeled in the absence of TdT were used as negative controls while cells pretreated with DNase I served as positive controls.
DNA electrophoresis. DNA extraction and electrophoresis were performed as described previously, with slight modifications (15). DNA extraction was performed with GenomicPrep Cells and a Tissue DNA Isolation kit (Amersham Pharmacia, Freiburg, Germany) according to the manufacturer‘s instructions. Up to 5 x 106 monocytes were first lysed by a hypotonic lysing buffer (10 mM Tris, 1 mM EDTA, and 0.2% Triton X-100, pH 7.4). After RNase treatment for 45 min at 37°C, protein precipitation was done by centrifugation by 16,000 x g for 3 min. Then DNA precipitation was performed by centrifugation with 70% ethanol followed by rehydration of DNA in hydration solution. Equal amounts of DNA were loaded on 1.5% agarose gel and separated by electrophoresis for 1 h at 80 V. The lower detection limit for visualization of oligonucleosomal bands was 1.0 µg of DNA.
Transmission electron microscopy (TEM)
Typical morphologic alterations indicative for apoptosis were evaluated by electron microscopy as described previously (15). Cells were washed off the culture plates, centrifuged, fixed in 1% glutaraldehyde/0.1 M sodium cacodylate-HCl (pH 7.4), and postfixed in 1% OsO4/0.15 M sodium cacodylate-HCl (pH 7.4). Samples were dehydrated in an ascending ethanol series and embedded in epoxy resin (Epon 812). Ultrathin sections were mounted on 150-mesh Formvar-coated copper grids and poststained with aqueous-saturated uranyl acetate and 2% lead citrate before being examined on a Philips CM 10 electron microscope (Philips Electronic Instruments, Mahwah, NJ) at an accelerating voltage of 60 kV.
Evaluation of cell necrosis
Viability of monocytes after different treatments was determined by trypan blue exclusion or PI uptake of nonpermeabilized cells using flow cytometry.
Flow cytometric caspase-3 and hsp-70 detection
Monocyte expression of activated caspase-3 was determined by flow cytometry. Monocytes were washed in PBS, fixed with 4% paraformaldehyde, and permeabilized with 0.1% saponin. For staining, a mAb against activated caspase-3 or hsp-70 (5 µg/ml, respectively) was applied for 20 min.
Detection, induction, and inhibition of phagocytosis
Phagocytosis of platelets was shown by TEM and specific membrane linking of platelets with a PKH67 Green Fluorescent Cell Linker kit (Sigma-Aldrich) according to the manufacturer’s instructions. In brief, platelets were resuspended in cell diluent and mixed with PKH67 dye in equal volumes at room temperature for 2 min. Staining reaction was blocked by adding 2 ml pure FCS for 1 min. After centrifugation of thrombocytes by 2000 x g for 10 min, they were washed extensively and resuspended in RPMI 1640 containing 0.2% FCS. Labeled platelets were used for coincubation experiments. Parallel detection of PI-stained nuclei (red fluorescence) and green fluorescence of labeled platelets by flow cytometry allowed recognition of apoptotic vs intact monocytes with and without ingested platelets. Latex beads (5 x 106/ml) or zymosan particles (10 x 106/ml) boiled for 30 min and opsonized with human AB serum were added to monocytes after 20 h and were washed off the cocultures after 4 h. For inhibition of phagocytosis, cytochalasin D (30 µM) was added to monocytes 30 min before adding platelets and washed off carefully after 1.5 h.
Semiquantitative RT-PCR
RNA isolation from monocytes after stimulation was conducted
using an RNeasy kit (Qiagen, Hilden, Germany) according to the
manufacturer’s instructions. Before transcribing into cDNA, DNase
(DNase I, RNase free; Boehringer Mannheim) digestion was performed.
cDNA was synthesized after the addition of 5 µM random primers
(pd(N)6; Roche Diagnostics, Mannheim, Germany), 1
mM dNTPs (New England Biolabs), and incubation at 37°C with Moloney
murine leukemia virus reverse transcriptase (Stratagene, Heidelberg,
Germany). Contamination with DNA was excluded by performing PCR from
templates incubated without reverse transcriptase. The primers used for
PCR amplification were 5'-ATG GAT GAT GAT ATC GCC GCG-3' and 5'-TCT CCA
TGT CGT CCC AGT TG-3' (human 
-actin, 248 bp), as well as 5'-CAC CAC
CTA CTC CGA CAA CCA-3' and 5'-GCC CCT AAT CTA CCT CCT CAA TG-3' (human
hsp70, 644 bp). The PCR mixture (40 µl) contained 2 mM
MgCl2, 0.2 mM dNTP, 1 µM primer, and 1 U
Taq DNA polymerase. Samples were amplified during 30 cycles
by 60-s denaturation at 94°C, 30 s annealing at 62°C (hsp70),
or 55°C (-actin) and 60-s elongation in a Peltier thermal cycler
(Biometra Uno II Thermocycler; Biometra, Göttingen, Germany).
For semiquantitative PCR, the relation between the expression of
-actin and hsp70 was analyzed. Signal intensity as measured by PCR
products was analyzed on a 1.5% agarose gel and visualized by ethidium
bromide staining. Densitometric quantification of PCR signals was
performed by the BioImage Intelligent Quantifier program (BioImage, Ann
Arbor, MI).
Western blotting
Cells were washed in PBS and resuspended at
106 cells/100 µl of sample buffer containing
2% SDS, 62.5 mM Tris-HCl (pH 6, 8), 10% glycerol, 5% 2-ME, and
bromphenol blue, heated at 95°C for 10 min, and stored at -20°C
until analysis. Samples, each containing 20 µg protein, were
separated by a NuPAGE Gel 4–12% BT (Invitrogen, Groningen, The
Netherlands) and transferred to nitrocellulose membranes (BA83 (0.2
µm); Schleicher & Schüll, Dassel, Germany) by a Western blot
modul (Invitrogen). Membranes were washed three times with
potassium phosphate buffer (0.05 M
K3PO4, pH 8.5) and
incubated with
digoxigenin-3-O-methylcarbonyl-
-aminocaproic
acid-N-hydroxysuccinimide ester) and Nonidet P-40 (0.01%
v/v) for 2 h. Then membranes were blocked with 5% fat-free milk
powder in TTBS buffer (0.01% Tween 20, 0.05 M Tris-HCl, and 0.15 M
NaCl, pH 7.5) and afterward incubated with the indicated Abs. Reactive
bands were visualized for detection of caspase-8 after incubation with
anti-rabbit IgG peroxidase-labeled Fab fragments, for caspase-9
after incubation with anti-mouse IgG peroxidase-labeled Fab
fragments and staining with BM Teton (Roche, Mannheim, Germany).
As an isotype-matched control for the primary Ab, mouse IgG2b was used.
BioImager master 3D was used for analysis of visualized bands.
Statistical analysis
Results are given as means ± SEM. For statistical analysis, the Mann-Whitney U test and for paired comparisons the Wilcoxon-signed rank test or Student’s t test was performed. A p < 0.05 was considered to be statistically significant. All experiments were performed at least five times with different buffy coats.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Coculture with platelets highly enhanced the life span of
monocytes. As shown in Fig. 1
A, 54 ± 5% of
monocytes were apoptotic when cultured for 60 h in medium
containing 0.2% FCS. Apoptosis was quantified by PI staining of
permeabilized cells. When platelets were added, apoptosis levels of
monocytes decreased in parallel with the number of added platelets and
reached at a monocyte:platelet ratio of 1:40 baseline levels of 12
± 4%. A coincubation time of 4 h was sufficient to exert this
protective effect. Similarly, when monocytes were rendered apoptotic by
a CD95 mAb (Fig. 1B), platelets reduced the monocyte
apoptosis level from 48 ± 7% to 25 ± 11%
(p < 0.05; monocyte:platelet ratio: 1:10).
This down-regulation of monocytic apoptosis by platelets was confirmed
when apoptotsis was determined by the TUNEL method (data not shown) or
by DNA electrophoresis. Fig. 2 shows that
internucleosomal DNA fragmentation of monocytes during constitutive
apoptosis (0.2% FCS) as well as anti-CD95-induced apoptosis was
highly reduced after the addition of platelets. Also, TEM revealed that
>50% of monocytes cultured in 0.2% FCS-containing medium for 60
h exerted nuclear and cytoplasmic alterations that were typical for
apoptosis. In Fig. 3A, three
intact monocytes after a culture in 5% FCS-containing medium are
depicted; in Fig. 3B, as a representative illustration, two
apoptotic monocytes after a culture in 0.2% FCS-containing medium are
shown. When monocyte morphology was studied after the coculture with
platelets, uptake of these cells and platelet ghosts into monocytic
lysosomes became visible (Fig. 3C). Despite the culture of
monocytes in 0.2% FCS-containing medium, <8% of studied monocytes
revealed nuclear alterations indicative for apoptosis. Trypsinization
of monocytes without consecutive disruption of platelets from monocytes
excluded plasma membrane invaginations and confirmed true phagolysosome
formation.
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We reasoned that phagocytosis of platelets might be the background
and prerequisite for enhanced monocyte survival. To confirm this, we
labeled platelets with a fluorescent linker to demonstrate thrombocyte
adherence and phagocytosis and studied the effects of the phagocytosis
inhibitor cytochalasin D (30 µM). As shown in Fig. 4, where apoptosis was detected by
reduced PI stainability of permeabilized monocytes, 54.7% of monocytes
were apoptotic after the culture in 0.2% FCS-containing medium for
60 h (upper left panel). Administration of
cytochalasin D for 1 h did not affect the viability of cells
(upper right panel). Addition of platelets exhibiting green
fluorescence caused a fluorescence shift to the right, indicating that
>70% of monocytes had ingested platelets or that adherence between
both the cell types took place. Low PI staining of apoptotic cells was
demonstrable in 16.2% (7.8 + 8.4%; Fig. 4, lower left
panel). When cytochalasin D was added, monocytes with ingested
platelets significantly decreased in number (Fig. 4, lower right
panel, upper right quadrant, 39.1%) and increasingly
became apoptotic (26.2 + 13.8%).
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We further tested whether phagocytosis per se was sufficient to enhance
the life span of monocytes and administered latex beads or zymosan
particles to monocytes. As shown by TEM for latex beads, a significant
number of monocytes became apoptotic in 0.2% FCS-containing
medium (>50%), although most monocytes had phagozytosed latex beads
(Fig. 3
D). In Fig. 5, it is
shown that neither latex beads nor zymosan particles influenced
monocyte apoptosis levels and lacked the protective effect shown for
platelets.
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Down-regulation of caspase-9 and -3 is involved in protection from growth factor-dependent apoptosis by platelets
Caspases-3 is one of the essential mediators of monocyte
programmed cell death (PCD). We looked for expression of active
caspase-3 by flow cytometry (16) with respect to
platelet-dependent down-regulation of monocyte apoptosis. Table I shows that constitutive apoptosis
induced by serum deprivation was characterized by a significant
expression of activated caspase-3, which was down-regulated to baseline
levels by addition of platelets. By studying the proximal regulatory
caspases-8 and -9 by immunoblotting, we could show that caspase-8 was
not activated by serum reduction and not affected by addition of
platelets (Fig. 6A). However,
when we performed Western blotting for caspase-9, a significant
reduction of pro-caspase-9 with the concomitant appearance of cleaved
fragments and subunits was demonstrable in monocytes under serum
starvation (Fig. 6B). Involvement and activation of
caspase-9 was reverted by the coculture with platelets. When apoptosis
was induced by CD95, caspase-3 stimulation occurred, but now platelets
could not block caspase-3 activation significantly (Table I).
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hsp-70 has been shown to exert antiapoptotic properties
(17). Thus, we determined hsp-70 expression on a mRNA
level by RT-PCR. Addition of platelets to monocytes highly increased
hsp-70-specific mRNA (Fig. 7).
Phagocytosis of latex beads could not stimulate hsp-70 expression. Heat
shock (47°C for 30 min) as a control led to an increase of hsp-70.
Serum starvation (0.2% FCS) significantly reduced hsp-70 expression as
compared with medium containing 5% FCS, underlining the importance of
hsp-70 for survival of monocytes. Similar results were obtained when
hsp-70 was detected at a protein level by flow cytometry (data not
shown).
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We further wanted to clarify whether activation of platelets had
any influence on their protective effects on monocytes. Platelets were
activated by preparation in hypotonic buffer and analyzed for CD62P
expression for mean fluorescent channel (MFC) using flow cytometry. As
shown in Fig. 8, reduction of apoptosis
of monocytes after addition of platelets was significantly dependent
from the grade of activation of platelets.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our data show that platelets are able to down-regulate the apoptosis program of cultured monocytes. This effect was drastic since already at a ratio between 1:20 and 1:40 (monocytes:platelets), which is exceeded by far in the blood, constitutive monocyte apoptosis induced by serum starvation was completely abrogated to baseline by suppressing caspase-9 and consecutively caspase-3. It has been shown by different groups that platelets themselves are equipped with a whole set of caspases, non-caspase proteinases, and substrates for these enzymes which all depend on the activation status of thrombocytes (19, 20, 21). It is conceivable that some accessories on the platelet side interfere with the counterparts on the monocyte side after phagocytosis and digestion. Since platelets down-regulated monocyte caspase-9 and -3, more a contribution of substrates rather than caspases by platelets for monocytic proapoptotic enzymes is supposed. Immunoblotting of platelet lysates excluded a significant amount of caspases supplied by these cells. Perhaps platelet compounds reactivate the energy household of monocytic mitochondria which is injured by serum deprivation (22, 23, 24). We could show that in the case of serum factor-dependent apoptosis platelets inhibited caspase-9 and the distal executive caspase-3. In the case of CD95-mediated PCD, platelets rescued monocytes from apoptosis without significant down-regulation of caspase-3. Our experiments could not completely elucidate at what stage platelets interfered here with the apoptosis pathway of monocytes. Probably, hsp are active and protective in this apoptotic pathway (25). We conclude that caspase-3 inhibition is not generally and probably only indirectly involved in the cytoprotective actions of platelets (26).
Monocyte apoptosis was studied and quantified by different methods,
such as flow cytometry using PI staining of nuclei after cell
permeabilization, TUNEL staining, DNA electrophoresis, or electron
microscopy, which all gave similar results. All of the nuclear
alterations indicative for PCD were suppressed by the coculture with
platelets. When we tried to determine monocyte apoptosis on the cell
membrane by annexin V staining, it soon became obvious that freshly
isolated platelets expressed significant amounts of phosphatidylserine
at their surface binding annexin V. This is in accordance with earlier
findings describing phosphatidylserine exposure of platelets as an
activation marker indicating procoagulant properties (27).
A recent study by Brown et al. (28) also demonstrated
annexin V binding by cultured platelets which increased by aging. It
was concluded that platelets cultured in vitro undergo constitutive
apoptosis by a caspase-independent pathway which was typically followed
by recognition and uptake by phagocytes. Taking these data into
account, we think that platelets mimic apoptotic cells and cross-talk
with monocytes/macrophages or other phagocytes in a fashion as
apoptotic cells do. In our study, platelets were fully viable as
determined by morphology (TEM), preserved expression of surface
markers, and ability of stimulation by thrombin. TEM demonstrated that
monocytes in culture rapidly captured platelets which first adhered at
the surface and consecutively were phagocytosed. Thus, intact platelets
may share properties of apoptotic cells and can be used to study the
interaction between apoptotic cells and phagocytes. We could show that
the uptake not merely adherence of platelets was the essential step for
the enhanced life expectancy of monocytes which was also paralleled by
hsp-70 induction. Inhibition of the uptake of thrombocytes by
cytochalasin D abrogated the protective effects of platelets. True
phagolysosome formation not merely platelet invagination was essential
for cytoprotection because trypsinization of monocytes had no influence
on ingestion rates. Adherence of platelets on the surface of monocytes
was insufficient for down-regulation of apoptosis, because short
coculture times of 1 h with consecutive washing off thrombocytes
not allowing significant uptake could not modulate cell senescence.
Platelet lysates were ineffective in suppressing monocyte PCD. In
contrast, phagocytosis of particles such as latex beads or zymosan did
not alter monocyte senescence, suggesting that the uptake process per
se is not sufficient to modulate PCD and that platelet surface
receptors or intracellular compounds released in phagolysosomes are
needed. Activation of platelets enhanced the protective, antiapoptotic
effects on monocytes. As the background and explanation for this,
activation of platelets induced higher uptake rates as detected by TEM.
Higher numbers of ingested platelets are able to provide bigger amounts
of protective compounds. The important conclusion can be drawn by our
data that phagocytosis of platelets by monocytes is not an inert
process leaving the phagocyte unaffected but evokes a whole cascade of
processes in the phagocyte shown here by the induction of survival
signals. Previously, it has been suggested that the disposal of
apoptotic cells was not accompanied by secretory or inflammatory
responses until we and others described that active and
mainly immunosuppressive pathways were evoked by apoptotic
cell phagocytosis, leading to enhanced production of anti-inflammatory
IL-10, TGF-, prostanoids or IL-6, or even to down-regulation of
proinflammatory cytokines (29, 30, 31). In our view, better
survival of the phagocyte due to platelet disposal fits in this scheme
of anti-inflammation since more surrounding cells in a vascular
lesion, including platelets or apoptotic inflammatory cells, can be
taken up and removed.
Further studies will elucidate which receptors on monocytes and platelets are primarily engaged in uptake and downstream processes and which signaling pathways are involved. Although aggregation between monocytes and platelets has been observed in the blood and called platelet satellitism, we do not think that this phenomenon is of major importance within the bloodstream under physiological conditions. Shear stress might disrupt the clots. However, in the vessel wall or at inflammatory sites when platelets are activated and the contact time is long enough these aggregates might become relevant.
In conclusion, our study shows that ingestion of platelets by monocytes suppresses their constitutive or induced apoptosis program primarily by affecting caspase-9, -3, and hsp-70 expression. These findings may have important implications for the understanding of progression and resolution of thrombotic and inflammatory vascular disorders or atherosclerosis and may offer therapeutic options.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Detlef Lang, Department of Medicine D, University Hospital Münster, Albert-Schweitzer-Strasse 33, D-48129 Münster, Germany. E-mail address: langd{at}uni-muenster.de
3 Abbreviations used in this paper: hsp, heat shock protein; PCD, programmed cell death; PI, propidium iodide; TEM, transmission electron microscopy; MFC, mean fluorescent channel.
Received for publication February 28, 2002. Accepted for publication April 11, 2002.
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, PGE2, and PAF. J. Clin. Invest. 101:890.[Medline]This article has been cited by other articles:
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M. C. Liebau, D. Lang, J. Bohm, N. Endlich, M. J. Bek, I. Witherden, P. W. Mathieson, M. A. Saleem, H. Pavenstadt, and K.-G. Fischer Functional expression of the renin-angiotensin system in human podocytes Am J Physiol Renal Physiol, March 1, 2006; 290(3): F710 - F719. [Abstract] [Full Text] [PDF]
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X174 HaeIII digest. Densitometric
determination of hsp-70-specific RT-PCR signals of differently treated
monocytes is given in B.