The JI
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     
 

The Journal of Immunology, 2007, 178: 6404-6415.
Copyright © 2007 by The American Association of Immunologists, Inc.
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
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 Similar articles in this journal
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 Google Scholar
Google Scholar
Right arrow Articles by Kadiu, I.
Right arrow Articles by Gendelman, H. E.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Kadiu, I.
Right arrow Articles by Gendelman, H. E.
Right arrowPubmed/NCBI databases
*Substance via MeSH

Cytoskeletal Protein Transformation in HIV-1-Infected Macrophage Giant Cells1

Irena Kadiu*,§, Mary Ricardo-Dukelow*,§, Pawel Ciborowski*,{dagger},§ and Howard E. Gendelman2,*,{ddagger},§

* Laboratory of Neuroregeneration, Department of Pharmacology and Experimental Neuroscience, {dagger} Department of Biochemistry and Molecular Biology, {ddagger} Department of Internal Medicine, and § Center for Neurovirology and Neurodegenerative Disorders, University of Nebraska Medical Center, Omaha, NE 68198


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The mechanisms linking HIV-1 replication, macrophage biology, and multinucleated giant cell formation are incompletely understood. With the advent of functional proteomics, the characterization, regulation, and transformation of HIV-1-infected macrophage-secreted proteins can be ascertained. To these ends, we performed proteomic analyses of culture fluids derived from HIV-1 infected monocyte-derived macrophages. Robust reorganization, phosphorylation, and exosomal secretion of the cytoskeletal proteins profilin 1 and actin were observed in conjunction with productive viral replication and giant cell formation. Actin and profilin 1 recruitment to the macrophage plasma membrane paralleled virus-induced cytopathicity, podosome formation, and cellular fusion. Poly-L-proline, an inhibitor of profilin 1-mediated actin polymerization, inhibited cytoskeletal transformations and suppressed, in part, progeny virion production. These data support the idea that actin and profilin 1 rearrangement along with exosomal secretion affect viral replication and cytopathicity. Such events favor the virus over the host cell and provide insights into macrophage defense mechanisms used to contain viral growth and how they may be affected during progressive HIV-1 infection.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Mononuclear phagocytes (MP;3 monocyte, dendritic cells, tissue macrophages, and microglia) are principal target cells and vehicles for HIV dissemination. Virus can persist within MP for prolonged time periods despite robust innate immune responses that include phagocytic, microbicidal, and secretory activities (1, 2). Common morphological and functional changes linked to productive viral replication include altered cell motility, Ag presentation, and cellular differentiation (3, 4). Such changes occur as monocytes leave the bloodstream and enter infected tissue where they undergo transformation to tissue macrophages and as a consequence of activation acquire proinflammatory secretory phenotypes (5). Such processes are typical of what occurs within the CNS during progressive HIV replication. Indeed, during advanced disease, monocyte-macrophages enter the brain within sites of inflammation and active viral replication and commonly form multinucleated giant cells (MGC) in attempts to contain infection. The formation of MGC occurs with profound astrogliosis, myelin pallor, and neuronal dysfunction. These are pathological signatures of HIV encephalitis and are linked to cognitive and motor deficits in susceptible people (6). The process of MGC formation is thought to occur as a consequence of interactions between the viral envelope glycoprotein, CD4 and other cell surface proteins (7, 8, 9, 10).

Despite more than two decades of research into the interactions between HIV and the macrophage, how profound changes in MP secretory, chemotactic, microbicidal, and Ag presentation activities during viral infection remain incompletely understood. Likely, such events affect MP responses to viral infection and a broad range of virus-induced inflammatory responses and are not specific for a single microbe (11). Indeed, morphological and functional cell changes elicited as a consequence of HIV infection are not virus specific and are operative during interactions with a broad range of infectious processes that include, but are not limited to, mycobacterial, fungal, protozoan, and bacterial pathogens (12). In all, the outcome of microbial-MP interactions serves as a means to control or eliminate pathogen growth while in many cases, simultaneously enhancing dissemination and subsequent tissue damage (13). Thus, elucidating the mechanisms underlying the molecular and biochemical changes in MP would have broader implications in infection and immunity.

Viruses as obligate cell parasites have evolved into manipulators of host cell functions. Accordingly, viruses often remodel the cytoskeleton of target cells to convert one of the cellular barriers to viral replication into an advantageous vehicle for the virus which facilitates the generation of infectious progeny virions (14). Surprisingly, little is known about the mechanisms used by HIV to exploit host cell cytoskeletal dynamics. However, HIV is known to polarize actin and control microtubule organization. This enables spread of virus from donor to target cells in close cell contacts termed "virological synapses" in T cells (15). We now have investigated the influence of HIV-1 infection for its ability to remodel cytoskeletal structures and affect macrophage MGC formation. In the course of proteomic analyses of the HIV-1-infected, monocyte-derived macrophage (MDM) secretome, we observed that both actin and profilin 1 were secreted in abundance and preceded macrophage cytopathology. Confocal microscopy analysis showed recruitment of profilin 1 and actin from the cytoplasm to the cellular membrane during the early stages of MGC and increased levels within the cytoplasm following cell fusion events. Most importantly, inhibitors of cytoskeletal rearrangements directly affect profilin 1-mediated actin stress filament/podosome formation, cell-cell contacts, and HIV replication. Based on these observations, we posit that actin and profilin 1 cellular transformation is triggered by HIV-1 infection. Modulations of the cytoskeleton are critical events required for the virus to complete its life cycle and lead to an attempt by the macrophage to contain ongoing viral growth.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Isolation, cultivation, and HIV-1 infection of human MDM

PBMC from HIV-1, HIV-2, and hepatitis seronegative human donors were obtained by leukophoresis in full compliance and approval of the University of Nebraska Medical Center Institution Review Board. Monocytes were purified by countercurrent centrifugal elutriation (16) then seeded onto T-75 or 250-ml Teflon flasks for adherent and suspension cultures, respectively. Cells were cultured in DMEM (Invitrogen Life Technologies) supplemented with 10% heat-inactivated pooled human serum, 1% glutamine, 10 µg/ml ciprofloxacin (Sigma-Aldrich), and 1000 U/ml highly purified M-CSF (a gift from Wyeth, Cambridge, MA) and maintained in a 37°C and 5% CO2 atmosphere. On days 2 and 5, half of the medium was exchanged. After seven days in culture, MDM were infected with HIV-1ADA (a macrophage tropic viral strain) at a multiplicity of infection (MOI) of 1, or left untreated (controls). On day 1 after infection, a full medium exchange was performed followed by a half medium exchange every 2–3 days thereafter in medium devoid of M-CSF. Medium was exchanged on day 2 after infection without phenol red. On days 3, 5, 7, 10, and 12 after infection, cells were exchanged with additive-free DMEM for an additional 24 h of incubation. Reverse transcriptase (RT) activity was determined in culture supernatant fluids as previously described (17).

Cell viability assays

Cell viability (mitochondrial activity) in control, HIV-1-infected, and poly-L-proline (P-L-P)-treated cells grown in adherence was measured by MTT reduction (18). In addition Live/Dead cell assay (Invitrogen Life Technologies) was used to assess cellular viability in adherent cultures at days 3, 5, 7, and 10 after infection in control, HIV-1 infected, and P-L-P-treated cells. Monocytes were cultured for 7 days in 90-well plates. Cells were treated with P-L-P 1 h before infection or left untreated and infected with HIV-1ADA at MOI of 1. At days 3, 5, 7, and 10 after infection, cells were rinsed with PBS and treated with 0.5 µM calcein AM and 4 µM ethidium homodimer-1 in PBS for 45 min, rinsed three times with PBS, followed by microplate centrifugations at 500 x g to avoid loss of dead cells by floatation. Cells permeabilized with 0.1% saponin were used as controls for both ethidium homodimer-1 and calcein AM. Live cells were distinguished from dead cells (red fluorescence; ex/em 495/635 nm) by the uptake of calcein AM to acquire a green fluorescence (ex/em 495/515 nm). Cell enumerations were performed using fluorescence microscopy and a M5 microplate fluorometer (Molecular Devices) (limit 1 ex/em 490/522 nm; limit 2 ex/em 530/645 nm) and normalized as the percentage of dead cells to uninfected controls.

Immunohistochemical tests

For determinations of numbers of HIV-1 p24 positive cells in adherent MDM, monocytes were grown in 4 well Lab-tech chamber slides (Invitrogen Life Technologies), infected with HIV-1ADA at a MOI of 1 or left uninfected. At days 3, 5, and 10 after infection cells were collected. MDM were stained with allophycocyanin-conjugated mouse anti-CD14 mAb (BD Biosciences) in 3% BSA in PBS for 30 min at room temperature. For HIV-1p24 staining and florescent microscopic examinations, cells were rinsed with PBS and fixed with 4% paraformaldehyde and 0.01% saponin (pH 7.4) for 1 h at room temperature, 50 mM NH4Cl to quench autofluorescence, and permeabilized in 100% methanol for 15 min at 4°C and 1% Nonidet P-40 (Sigma-Aldrich). Cells were incubated with FITC-conjugated mouse anti-HIV-1 p24 in 3% BSA (KC57 FITC; Beckman Coulter) for 45 min. A NIKON E-800 fluorescent microscope (Nikon) was used to evaluate HIV-1 p24-positive cells.

For suspension cell analyses, MDM were stained with allophycocyanin-conjugated anti-CD14 mAb (BD Biosciences) and FITC-conjugated IgG2a mouse anti-HIV-1 p24 (KC57 FITC; Beckman Coulter) as described above, and data acquisition was performed on a FACSCalibur flow cytometer (BD Biosciences) using CellQuest software (BD Biosciences). Live cells were gated by forward and side scatter using FCS Express V3 software (De Novo Software) (see Fig. 1). Data are shown as density plots of uninfected and infected cells at days 3, 5, and 10 after infection.


Figure 1
View larger version (28K):
[in this window]
[in a new window]

  		FIGURE 1. HIV-1ADA replication in MDM. MDM were infected with HIV-1ADA at a MOI of 1 after 7 days in suspension (A) or adherent (B) cultures. A, HIV-1 p24 and CD14 expression in control and infected human MDM. Cells were collected days 3, 5, and 10 after infection and stained with anti-CD14 allophycocyanin-conjugated (extracellular) and anti-HIV-1 p24 FITC-conjugated Ab (intracellular). FACS analysis determined in percentage of virus-infected MDM. B, HIV-1 p24 expression was assayed in infected MDM grown in adherence. HIV-1-infected cells were stained against CD14 and HIV-1 p24 (as above) at days 3, 5, and 10 after infection and counted by fluorescent microscopy. Levels of infection are presented as the ratio of HIV-1 p24 to the overall CD14+ population. Results are representative of experiments with four separate donors (n = 4; error bars = ± SEM).

 
For confocal microscopy examinations, cells were stained with mouse anti-profilin 1 mAb (clone 2H11; Synaptic Systems) and FITC-conjugated (goat) anti-mouse IgG mAb (Invitrogen Life Technologies), rhodamine-conjugated phalloidin (Invitrogen Life Technologies) to detect F-actin, and To-PRO as a nuclear stain for 3 h, 30 min, and 20 min, respectively. Each treatment was followed by triplicate 5-min washes with 0.1% saponin in PBS. Cells analyzed for HIV-1 p24 and profilin 1 or F-actin intracellular localization were stained with FITC-conjugated mouse anti-HIV-1 p24 Ab (KC57 FITC; Beckman Coulter), goat anti-profilin 1 mAb (Alexis Pharmaceuticals) and Alexa 598-conjugated (sheep) anti-goat mAb (Invitrogen Life Technologies), and rhodamine-conjugated falloidin (Invitrogen Life Technologies), respectively. Cells were mounted using anti-fade Pro-Long Gold mounting reagent (Invitrogen Life Technologies) and examined with a Zeiss LSM 510 META NLO microscope (Zeiss MicroImaging).

Culture supernatant and cell lysate preparations

On days 3, 7, 10, and 12 after viral infection, culture fluids and MDM from infected and control cultures were centrifuged for 10 min at 2000 x g and treated with lysis/deactivation buffer (0.1% Triton X-100, 50 mM Tris-HCl, and protease inhibitor mixture (Sigma-Aldrich)). The supernatants were concentrated using Amicon Centriplus centrifugal filter devices (Millipore) with a membrane cutoff of 5000 Da molecular mass and dialyzed for 18 h against water at 4°C using QuixSep microdialyzers (Membrane Filtration Products). Protein concentrations were determined using Bio-Rad DC Protein Assay (Bio-Rad).

Virus and vesicle preparations

Infected and control MDM culture fluids were pooled, clarified (10 min at 2,000 x g), filtered through 450- and 200-nm pore-size sterile filter (Beckman Coulter), and concentrated using Amicon centrifugal filter devices (Millipore) with a 100,000 molecular mass membrane cutoff. Fluids were depleted of vesicles using anti-CD45 paramagnetic microbeads (Miltenyi Biotec) as described previously (19). Briefly, culture fluids were treated with 2 µl/ml anti-CD45 microbeads for 1 h at room temperature with shaking and then placed in magnetic separators (Invitrogen Life Technologies) at 4°C for 20 h. The vesicles bound to microbeads were recovered and retained for further analysis, and supernatants were layered on 20% (w/v) sucrose-PBS cushion and centrifuged for 4 h at 120,000 x g at 4°C.

Surface-enhanced laser desorption and ionization time-of-flight (SELDI-TOF)

The protein profiles of culture supernatants were analyzed by SELDI-TOF ProteinChip assays (Ciphergen Biosystems). The chip types tested included weak cation exchange (WCX2), strong anion exchange (SAX2), normal phase (NP20), and immobilized metal affinity capture (IMAC30). Based on sensitivity and specificity of profile predetermination, the WCX2 ProteinChip was selected to profile the culture supernatants. The ionized proteins and their molecular mass/charge (m/z) ratios were assessed by SELDI-TOF as described previously (20). Spectra were collected and analyzed using PBS II ProteinChip Biosystems (Ciphergen Biosystems). All tests were performed in triplicate, in three independent experiments, and from three separate cell donors. The ProteinChip Reader was externally calibrated for each analysis for both low and high molecular mass regions using standard proteins. Peaks were automatically detected with Biomarker Wizard for ProteinChip software 3.2. Parameters for peak detection were first pass signal/noise (S/N) ratio 5, second pass S/N ratio 2, and mass tolerance 0.5%.

For the profilin 1 immunodepletion, 200 µg of protein from culture fluids was incubated with 20 µg of anti-profilin 1 rabbit polyclonal Ab for 24 h at 4°C (Novus Biologicals), followed by a 2-h incubation at room temperature with 100 µl of protein A/G agarose gel slurry (Pierce). Following incubation, the agarose beads were precipitated by centrifugation at 1000 x g for 5 min, and 5 µg of protein was collected from the profilin 1-immunodepleted fluids. Affinity-purified crystallography-grade profilin 1 (provided by Dr. G. Borgstahl, Eppley Institute, University of Nebraska Medical Center, Omaha, NE) and nondepleted culture fluids were used as positive controls.

One-dimensional SDS-PAGE

One-dimensional SDS-PAGE separation used a NOVEX (Invitrogen Life Technologies) system and 4–12% gradient gel. For protein identification by liquid chromatography mass spectroscopy (LC/MS-MS), 20 µg of protein was loaded per lane, and protein bands were detected using a Typhoon 9410 PhosphorImager (Amersham Biosciences-GE Health Care). Gel pieces excised from one-dimensional were cut into 2- to 4-mm fragments and destained in 200 µl of 50% acetonitrile and 50 mM NH4HCO3 at room temperature for 1 h, then dried, and 2 µl of 0.5% trypsin (Promega) and 50 µl of 10 mM NH4HCO3 were added to each sample overnight at 37°C. Peptides were extracted by washing gel pieces twice with 0.1% trifluoroacetic acid and 60% acetonitrile. Samples were transferred to vials, dried, and resuspended in 15 µl of water with 0.1% formic acid for automated injections.

Nanoelectrospray ionization-LC/MS-MS

Sequencing was performed using ProteomeX LC-MS/MS system (ThermoElectron) in nanospray configuration from bands excised from one-dimensional SDS-PAGE and loaded onto a microcapillary C18 column. Peptides were eluted and sprayed directly into the mass spectrometer using a gradient of water/acetonitrile and 0.1% formic acid. The spectra obtained were searched against the NCBI.fasta protein database using Bioworks 3.1 SR1 software provided by ThermoElectron.

Western blot assays

For these assays, 10 µg of protein from the cell lysates and 20 µg from the culture fluids were loaded per lane for one-dimensional gel electrophoresis. Human platelet protein (Novus Biologicals) was used as a positive control. Blots of samples from culture supernatants, vesicles, virus, and cell lysates were probed with rabbit anti-profilin 1 and anti-actin Abs (Novus Biologicals) and HRP-conjugated anti-rabbit IgG (Novus Biologicals). HIV-1 p24 from blots of viral extracts was detected using mouse mAb anti-p24 and HRP-conjugated polyclonal (goat) anti-mouse IgG (Novus Biologicals). Vimentin was detected using mouse anti-vimentin Ab and HRP-conjugated polyclonal (goat) anti-mouse IgG (Novus Biologicals). Signal intensities were determined by densitometry analysis using Gel-Expert software (Nucleotech).

Cytoskeletal inhibitors

MDM were treated with P-L-P to assess profilin 1 function. Before infection, cells were treated for 1 h with 3 µM solution of P-L-P or left untreated as controls. The optimal concentration for P-L-P was determined from previous dose-response experiments and assessing cell viability and function in MDM cultures for toxic effects. Following incubation, cells were rinsed with PBS and infected with HIV-1ADA at a MOI of 1. Supernatants and cells were collected for analysis by Western blotting, confocal, and cell viability assays. The Giant Cell Index (GCI; numbers of nuclei per cell) and the percentage of giant cells (ratio of cell number containing more than three nuclei to the overall cell population) were determined. MDM were plated for 7 days in 4-well Lab-Tek chamber slides (Invitrogen Life Technologies) at a concentration of 5 x 105 cells/well with or without P-L-P and virus infected at a MOI of 1. At days 3, 5, 7, and 10 after infection, slides were fixed and stained with Giemsa as described by the manufacturer (Invitrogen Life Technologies).

Protein modifications

Pooled protein (200 µg) from culture fluids of infected cells collected at days 3 and 5 after infection was incubated with 20 µl of 0.1 µg/µl mouse anti-phosphotyrosine mAb (Invitrogen Life Technologies), followed by a 2-h incubation at room temperature with 100 µl of protein A/G gel slurry (Pierce). Sample was supplemented with 500 µl of BupH TBS (Pierce) immunoprecipitation buffer centrifuged at 2500 x g for 5 min, and the agarose beads were precipitated. Supernatant was recovered and retained for immunoblotting. The pellet was treated with 100 µl of IgG elution buffer (Pierce) to elute the immune complex. Sample was centrifuged at 2500 x g for 5 min, and the supernatant was retained. Sample fractions and nontreated culture fluids were used for immunoblotting and stained against profilin1 and actin using rabbit anti-profilin 1, anti-actin, and HRP-conjugated anti-rabbit polyclonal Abs (Novus Biologicals).

Statistical analyses

All resulting data were analyzed for statistical significance by both Student’s t tests and a one-way ANOVA with Newman-Keuls posttest.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
HIV-1ADA MDM infection

To provide a biological framework for subsequent proteomic analyses, we first investigated both replication and cytopathic effects of HIV-1ADA infection in MDM of cells grown in suspension (Fig. 1A) and as adherent cultures (Fig. 1B). Measurements included the percentage of cells positive for HIV-1 p24 at days 3, 5, and 10 after viral exposure at an MOI of 1. Progeny virion production was measured by RT activity in culture fluids, and GCI and fusion activity were examined by morphology. These findings paralleled one another (see Fig. 7, A–C) and allowed correlations between measures of HIV-1 replication and cytopathicity with levels of secreted cytoskeletal proteins (see below).


Figure 7
View larger version (38K):
[in this window]
[in a new window]

  		FIGURE 7. Cytokeletal inhibitors affect HIV-1 replication and MDM fusion. Monocytes were propagated as adherent cells for 7 days, then treated for 1 h with 3 µM P-L-P before infection with HIV-1ADA at a MOI of 1. HIV-1-infected and -uninfected MDM without or with P-L-P served as controls. A, Profilin 1 inhibitor P-L-P suppresses productive infection. P-L-P significantly reduces progeny virus formation as shown by levels of RT activity. B and C, P-L-P suppresses cellular fusion. To generate indices of cell fusion, control and treated cells were stained with Giemsa, and numbers of nuclei per cell were counted for GCI measurements (B). Fusion activity was presented as a ratio (in %) of number of cells containing more than three nuclei to the total cell counts (C). D and E, Cell viability and P-L-P toxicity. To determine whether secretion of cytoskeletal proteins in the culture fluids could be due to cell death, both MTT (D) and live dead assays (E) were performed. On days 3 and 5 when maximal secretion of actin and profilin 1 were observed, cell death was limited. P-L-P showed minimal cytotoxicity. F, Disruption of actin filament formation and fusion. P-L-P-treated infected MDM were harvested on day 5. Cells were stained for profilin 1 (green), F-actin (red), and nuclei (blue) as described in Fig. 5 and analyzed by confocal microscopy. Panels show lack of actin filaments, disruption of profilin 1 recruitment to the plasma membrane, and no fusion activity. Arrows indicate G-actin aggregates randomly distributed throughout the cytoplasm. G, P-L-P reduces profilin 1 and actin secretion. Western blot analysis of culture fluids from untreated and P-L-P-treated, HIV-1-infected MDM against profilin 1 and actin revealed a reduction of profilin 1 and actin in the P-L-P-treated cells. Images were obtained at x100. Bars, 25 µm. (*, p < 0.05 and **, p < 0.001; error bars, ± SEM, n = 4).

 
SELDI-TOF fingerprints

To determine optimal protein binding conditions and spectral accuracy, the affinities of culture fluid proteins to chromatographic surfaces were tested. Based on these analyses, the WCX2 was shown to be most sensitive and specific and thus used for SELDI-TOF screening of MDM-secreted proteins. SELDI-TOF spectra of culture fluids at days 7, 9, and 12 after infection were obtained within a 5- to 150-kDa (m/z) range. This revealed 21 differentially expressed (up- or down-regulated or uniquely expressed) peaks among the infected and uninfected MDM culture fluids. Representative spectra of culture fluids at day 7 after infection in the 10- to 20-kDa region are shown in Fig. 2A. Arrows indicate 9 of 21 differentially expressed proteins in infected MDM vs the controls. An enlarged version of the marked peak at the 15- to 16-kDa range in Fig. 2A is shown in Fig. 2B. Arrow indicates one differentially expressed peak in the HIV-1-infected MDM culture fluids. Comparison of the molecular mass against the Swiss-Prot protein database matched profilin 1 as the "putative" protein to the up-regulated ~15-kDa peak. Spectral analysis of the 40- to 50-kDa region revealed a 43-kDa peak correlating with increased levels of actin (data not shown). To determine whether the 15-kDa peak was indeed profilin 1, HIV-1-infected fluids were immunodepleted with Abs to profilin 1 and spotted on a WCX2 chip. Moreover, parallel spectra were obtained with affinity-purified profilin 1 and crude culture fluids in which profilin 1 was not depleted (Fig. 2C). The absence of the 15-kDa peak in the profilin 1-immunodepleted culture fluids provided the first confirmation for the identity of profilin 1 and its up-regulation in HIV-1-infected MDM cultures.


Figure 2
View larger version (24K):
[in this window]
[in a new window]

  		FIGURE 2. SELDI-TOF analysis of HIV-1-infected MDM culture fluids shows profilin 1 secretion. MDM were infected with HIV-1ADA at a MOI of 1 after 7 days in culture. A, Representative spectra of control (top) and HIV-1-infected (bottom) culture fluids are shown within the 10- to 20-kDa range. Arrows indicate nine differentially expressed proteins. B, Representative spectra on culture fluids of control (dashed line) and HIV-1 infected (thick line) obtained by SELDI-TOF analysis on day 7 after infection using a WCX2 chip. Arrow indicates putative peak for profilin 1, which is present in HIV-1-infected MDM culture fluids. C, Spectral peak is identified as profilin 1. Top panel, SELDI-TOF profile of affinity-purified profilin 1. Spectra of the crude culture fluids (*CF) collected from HIV-1-infected MDM at day 5 after infection, and immunodepleted fluids for profilin 1 are shown in the middle and bottom panels, respectively. Spectra shown are representative of data obtained from four different donors.

 
Cytoskeletal proteins

To provide verification of our SELDI-TOF analyses, replicate HIV-1 and control MDM culture supernatants were fractionated by one-dimensional SDS-PAGE. The band fragments, including those corresponding to the molecular mass of 15 and 43 kDa, were excised, digested, and sequenced by LC-MS/MS. The tandem mass spectra were compared against spectra of the European Molecular Biology Laboratory (EMBL) nonredundant human protein database by using a SEQUEST search program. After filtering the results based on cross-correlation Xcorr (cutoffs of 2.0 for [M + H]1+, 2.5 for [M + 2H]2+, and 3.0 for [M + 3H]3+), peptides with scores >3000 and meeting {delta} cross-correlation scores ({Delta}Cn) >0.3 and fragment ion numbers >60% were deemed valid by these SEQUEST criteria thresholds. These were determined to afford >95% confidence for peptide identification (21). The identities of sequenced proteins were confirmed in samples obtained from three independent experiments. The peptide identifications were segregated into cytoskeletal and other proteins. The identified cytoskeletal proteins, including actin and profilin 1, which met the selection criteria are shown in Table I. Other proteins identified were reported elsewhere (22). In agreement with the SELDI-TOF results, profilin 1 and actin were identified. Quantitative immunoblots were subsequently required for verification and substantiation of differential expression.


View this table:
[in this window]
[in a new window]

  		Table I. Cytoskeletal proteins identified in HIV-1-infected MDM culture fluids

 
Quantitation of actin and profilin 1

To quantify cytoskeletal proteins in culture fluids, Western blot analyses for profilin 1 and actin in HIV-1 infected and control MDM cell lysates and culture fluids were performed (Fig. 3, A and B). Intracellular profilin 1 and actin levels were unchanged throughout the course of viral infection. In contrast, these proteins were found increased in culture fluids of HIV-1-infected MDM through day 7 after viral exposure. Interestingly, levels of profilin 1 and actin were decreased by days 9 and 12, corresponding to maximum MGC activity and cytopathicity. The increases in profilin 1 and actin secretion at days 5 and 7 suggest close associations between cytoskeletal secretion and the early stages of virus-induced MDM fusion events and were not related to cell death.


Figure 3
View larger version (52K):
[in this window]
[in a new window]

  		FIGURE 3. HIV-1-infected MDM secrete profilin 1 and actin. Cells and secreted fluids collected from control (–) and HIV-1-infected MDM (+) were examined from days 5 to 12. A, Intracellular profilin 1 and actin expression throughout infection. Western blot analyses for both profilin 1 and actin in cell lysates revealed no changes between infected and control MDM in the course of infection. B, Profilin 1 and actin are secreted. Immunoblot assays for profilin 1, actin, and vimentin on culture fluids collected at parallel time points showed increased levels of both of these proteins from virus-infected MDM as compared with controls. C, Secreted profilin 1 and actin are phosphorylated. Immunoprecipitation using anti-phosphotyrosine mAb, followed by immunoblotting of unaltered (control) and phosphotyrosine-depleted culture fluids, as well as immunoprecipitate (IP) against profilin 1 and actin, shows these proteins are tyrosine phosphorylated. D, Profilin 1 and actin packed inside exosomes. Immunoblotting against profilin 1 and actin of CD45+ vesicles isolated from pooled culture fluids of infected MDM at day 3, 5, and 7 postinfection demonstrates presence of these proteins in the CD45+ vesicle fraction. E, Thin section transmission electron microscopy imaging of isolated progeny HIV-1 non-CD45 depleted (control; arrows indicate vesicles), recovered CD45+ vesicles, and HIV-1 depleted of CD45+ vesicles. Images show each fraction is organelle free and that CD45 depletion of vesicles from the culture fluids is efficient. Bars, 500 and 100 nm; images were obtained at x31,000 and x153,000, respectively.

 
To preclude presence of these cytoskeletal proteins in the culture fluids as a result of dead or dying cells, we monitored, in parallel, cell viability in infected and control MDM grown in adhesion by performing live/dead cell assays at these time points (see Fig. 7E). Cells were stained with calcein AM (live cells-stained green) and Eth-D1 (dead cells stained red) and read on a fluorescent microplate reader. Readings from cells used as positive and negative controls for each dye were included in calculating the percentage of dead cells using the formula provided by the manufacturer. Live/dead cell assays revealed a low cell death activity compared with the uninfected cells at days 3 and 5, which mark maximal levels of extracellular profilin 1 and actin. These data, taken together, provide clear evidence that HIV-1-induced actin skeleton transformation can affect the viral infection and cellular fusion and that the secretion of cytoskeletal proteins are not as a consequence of cell death or injury. MTT assays supported these findings (see Fig. 7D).

Next, we evaluated whether the secreted profilin 1 and actin were posttranslationally modified. MALDI-TOF and LC/MS-MS of trypsin digested actin and profilin revealed phosphorylation activity on amino acid tyrosine (data not shown). Immunoprecipitation assays using Abs to phosphorylated tyrosine performed on culture fluids from infected MDM and followed by Western blot assays for profilin 1 and actin showed that both proteins were in fact tyrosine phosphorylated (Fig. 3C). These findings, together with previous reports regarding vimentin, actin, and profilin 1 secretion by MDM (19, 23), support the presence of phosphorylated actin and profilin 1 proteins in the culture fluids as a result of infection and giant cell formation.

Profilin 1 and actin exosomal transport

We next investigated the mode of extracellular transport of profilin 1 and actin. Based on previous studies demonstrating secreted proteins may be packed within distinct vesicle subtypes expressing the CD45 receptor (24, 25, 26), we hypothesized that the cytoskeletal proteins could be either secreted within exosomes or packaged with progeny virus. We isolated progeny virions and CD45+ vesicles from HIV-1-infected MDM culture fluids. Fluids pooled in the course of viral infection were cleared from cellular debris by centrifugation followed by two filtrations through 400- and 200-nm filters, immunodepleted of CD45-expressing vesicles and isolated as pellets following ultracentrifugation on a 20% sucrose cushion. Western blot tests performed on virion-enriched fractions and recovered CD45+ vesicles demonstrated that actin and profilin 1 were not packed within progeny virions; however, they were readily detected in CD45+ vesicles (Fig. 3D). The purity of progeny virus and CD45+ fractions were confirmed by RT activity, HIV-1 p24-specific immunoblots (data not shown), and thin sectioning of the pellet by transmission electron microscopy (Fig. 3E).

Viral replication and actin and profilin 1 secretion

To assess the dynamics of actin and profilin 1 secretions and its relationship to progeny virion production and MGC formation, serial collections of culture fluids were performed during the early and later stages of viral infection. MDM were infected with HIV-1ADA at a MOI of 1. Cells were washed with serum-free medium, and culture fluids were sampled every 8 h for 24 h on days 3, 5, 10, and 12 after viral exposure. After the last sampling at 24 h, cells were maintained in serum containing medium until the next collection. Serum deprivation during sampling was performed to 1) enhance the outcome of infection causing cell stress by serum starvation, 2) avoid spurious results because the gene that codes for actin contains a serum response element and could be induced as such through engagement of its serum response factor (27), and 3) to maximize proteomic analyses as high albumin content in the supernatants masks the presence of low abundance proteins and limits their detection. Quantitative assessment for profilin 1 and actin by Western blot assays and RT were performed simultaneously. RT and densitometry data shown were previously normalized to values obtained by similar analysis on uninfected controls. Fig. 4 compares dynamics of profilin 1 and actin secretion and progeny virion production. Interestingly, virus production preceded profilin 1 release into the culture fluids (Fig. 4A). The dynamics of actin secretion were shifted, yet similar, to that observed for profilin 1 (Fig. 4B). Western blots analyzed by densitometry are shown in the panels below A and B of Fig. 4. These results support distinct patterns of profilin 1 and secretion in the course of infection, as well as the basis for protein-virus interactions in regards to critical cellular response to HIV-1 infection.


Figure 4
View larger version (26K):
[in this window]
[in a new window]

  		FIGURE 4. Dynamics of actin and profilin 1 protein and progeny virion secretion. MDM were infected with HIV-1ADA at a MOI of 1 after 7 days in culture. Infected and control MDM were incubated in serum-free medium, and culture fluids were collected in three 8-h intervals (collection points) starting at day 3 after infection. A and B, Dynamics of profilin 1 and actin secretion and progeny virion production. RT assays were performed parallel to densitometry analysis (intensity of band/area) on Western blots of HIV-1-infected MDM culture fluids for profilin 1 (A, bottom panel) and actin (B, bottom panel) and normalized to uninfected control of similar time points.

 
Cytoskeletal transformation and giant cell formation

Based on the data obtained and the dynamics of protein secretion, we reasoned that profilin 1 and actin secretion may be due to actin cytoskeleton reorganization by HIV-1. At days 5 and 10 after viral infection, MDM were fixed and stained for profilin 1 (Fig. 5, A, D, G, J, and M) and F-actin (Fig. 5, B, E, H, K, and N), and images were merged (Fig. 5, C, F, I, L, and O). Confocal images of MDM grown in suspension revealed massive profilin 1 and F-actin reorganization and recruitment to the cell membrane in association with fusion events in HIV-1-infected MDM (Fig. 5, G–I; arrows indicate profilin 1 and actin localization at the fusion interface). Unlike the infected MDM, images of the uninfected cells at a similar time point (day 5) revealed profilin 1 and actin were distributed homogeneously throughout the cytoplasm (Fig. 5, A–C).


Figure 5
View larger version (88K):
[in this window]
[in a new window]

  		FIGURE 5. Profilin 1 and actin distribution in HIV-1-infected MDM. Human MDM were propagated as suspension (G–O) or adherent cultures (A–F) for 7 days then infected with HIV-1ADA at a MOI of 1. Uninfected (A–C and J–L) and infected (D–F, G–I, and M–O) MDM were collected on days 5 (A–I) and 10 (J–O) after infection. MDM grown in suspension were sedimented onto slides before analysis. Cells were stained against profilin 1 with mouse anti-profilin-1 (mAb) and FITC-conjugated anti-mouse IgG (green), F-actin by rhodamine-conjugated phalloidin (red), and nuclei by To-PRO (blue). Images in G–I indicate increased recruitment of profilin 1 and actin to the contact interface between infected MDM at the initiation of cellular fusion (indicated by arrows). Profilin 1 and actin colocalization in a fully formed MGC at day 10 is shown in M–O. D–F, Profilin 1 and F-actin colocalization in the podosomes of infected MDM at day 5 postinfection in adherent cells. Arrows indicate profilin 1-mediated actin podosome formation and establishment of intercellular contacts between cells. Images reveal newly reorganized profilin 1 and actin from the cortical region of the cellular membrane to the perinuclear region of MGC (indicated by arrows). The merged images are presented in differential interference contrast (DIC). Images were obtained at x100. Bars, 25 µm.

 
At the latter stages of infection (day 10) and completion of cellular fusion (~100 nuclei/cell), profilin 1 and actin showed reorganization from the membrane back to the cytoplasm and more distinctly localized in the perinuclear region as indicated by the arrows (Fig. 5, M–O), whereas the uninfected cells maintained a homogeneous distribution of actin and profilin 1 (Fig. 5, J–L). In replicate experiments performed on adherent MDM, images revealed profilin 1-mediated actin stress filament formation at the active edges of the membrane in the form of podosome-like extensions and establishment of focal contacts with the adjacent cells as early as day 5 after infection (Fig. 5, D–F). These results support the notion that profilin 1 and actin translocation to the plasma membrane, as well as podosome formation, are necessary events preceding formation of cell-cell contacts and intercellular membrane fusion.

Actin and profilin 1 transformation, viral replication, cell fusion, and cytopathicity

Confocal analysis performed on days 5 (Fig. 6, A–F) and 10 (Fig. 6, G–L) after viral infection revealed significant colocalization of HIV-1 p24 with profilin 1 (Fig. 6, A–C and G–I) and actin (Fig. 6, D–F and J–L). This was seen at day 5 in MDM throughout the length of cell processes with intense staining at the area of focal contact with the next cell (as indicated by arrows in Fig. 6, C and D). At the completion of cellular fusion at day 10, HIV-1 was colocalized with profilin 1 and actin in the perinuclear region of the MGC (Fig. 6, G–I and J–L). Fig. 5, M–O, showed a similar pattern of profilin 1 and actin colocalization in the perinuclear region of the giant cells. These data suggest that HIV-1 may use the actin cytoskeleton intercellular network to increase the efficiency of infection and cellular fusion.


Figure 6
View larger version (59K):
[in this window]
[in a new window]

  		FIGURE 6. Profilin 1, actin, and HIV-1 distribution in infected MDM. Human MDM were propagated as adherent cultures, then infected with HIV-1ADA at a MOI of 1, and cells were immunostained at days 5 (A–C) and 10 (G–I) for profilin 1 with goat-anti-profilin mAb and anti-goat Alexa 533 (red). For HIV-1 detections, FITC-conjugated anti-HIV-1 p24 (green) were used. Cells at day 5 (D–F) and 10 (J–L) were stained for F-actin and HIV-1p24 with rhodamine-phalloidine (red) and FITC-conjugated anti-HIV-1 p24 (green). To-PRO (blue) was used as a nuclear stain. Images demonstrate colocalization of HIV-1 with cytoskeletal proteins during MDM fusion. Robust HIV-1 staining is noted along the actin stress filaments (arrows; A–F) and at the point of contact between the podosome from one cell and the cell membrane of the next (shown by top arrow A–C). At the completion of fusion (day 10; G–L), images reveal a distinct colocalization of HIV, profilin 1, and F-actin in the perinuclear region. Images were obtained at x60. Bars, 50 µm.

 
After day 7 in culture, uninfected MDM were treated with 3 µM P-L-P. Following 1-h incubation, cells were infected with HIV-1ADA at a MOI of 1. The inhibitor for profilin 1, P-L-P, binds with high affinity to the phosphorylated form of profilin 1 and suppresses its interaction with G-actin (monomeric form) and consequently the process of actin filament formation (28, 29, 30). Escalating concentrations of these drugs were tested on both HIV-1-infected and control MDM to determine an optimal therapeutic index (drug efficacy with limited toxicity; Fig. 7E). Cell viability assays showed treatment with 3 µM solution of P-L-P exerted maximal suppression of viral progeny production (Fig. 7A) and cell fusion (Fig. 7, B and C) with minimal associated cytotoxicity (Fig. 7, D–F). Suppression of productive infection along with cellular fusion appears to be a direct reaction to the disruption of actin filament formation by P-L-P because phagocytic activity and reactive oxygen species production were not affected in inhibitor-treated MDM (data not shown). Along with significantly (*, p < 0.05 and **, p < 0.01) reduced RT activities at days 3–10 postviral exposure, MDM treated with P-L-P showed complete disruption of the F-actin intercellular network and reduced profilin 1 and actin secretion (Fig. 7, F and G). These data suggest HIV-1-induced actin and profilin 1 alterations and reorganization play important roles in productive infection and cellular fusion.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Links between HIV-1-induced cytoskeletal alterations, productive viral replication, and membrane fusion were shown using combinations of proteomics and immunologic approaches. HIV-1 infection of MDM induced polarized recruitment of actin and profilin 1 to the plasma membrane, secretion through exosomes, induction of podosome-mediated intercellular contacts, and giant cell formation. These underlie cellular events necessary for productive HIV-1 replication as inhibition of profilin 1 with P-L-P disrupts profilin 1 transformation suppresses progeny virion production and cell fusion.

Studies by others investigating the interactions between cytoskeletal proteins and HIV-1 have shown virus-induced, actin-dependent receptor colocalization regulates viral entry, podosome formation (9), and intercellular virion transfer across a "virological synapse" (15). In addition, actin reorganization induced by HIV-1/coreceptor interactions and Rac-1 GTPase are events demonstrated to precede cellular fusion and MGC formation (31). Investigations in various model systems have shown profilin 1 to be a common denominator in the intimate cross-talk of small GTPase (Rac, {rho}, and Cdc42)-mediated polymerization of actin (32, 33), receptor activities, guidance of focal adhesions (34, 35), and membrane trafficking (36, 37, 38, 39). Because focal adhesions and stress filament formation is often an artifact of cell growth on adhesive substrate, we performed replicate experiments of MDM infection in suspension and observed a consistent and similar pattern of actin and profilin 1-polarized recruitment at the fusion interface of adjacent cells.

Other intercellular pathogens such as Listeria monocytogenes (40, 41) and Shigella flexneri (42) use profilin 1 and actin to affect its propulsion across the cytoplasm and spread into neighboring cells without entering the extracellular space. Moreover, enveloped viruses use the cortical actin network as a scaffold to orchestrate the lateral mobility of the virion-receptor complex in the plane of the plasma membrane, as well as their dissemination across cells (43). Viruses use the actin stress filaments to engage the surface of the cell on lipid rafts in search of internalization or endocytic sites following receptor binding (44, 45). This is likely facilitated by coupling of the virus through the cytosolic domain of its receptor to an actin filament at the base of the filopodium (46, 47). In the final stages of the viral life cycle, progeny virions budding from the host’s cytoplasm use pseudopodium-mediated intercellular contacts as fast tracks for viral dissemination. This process is often associated with up-regulation of vesicle recycling to augment the plasma membrane surface area and facilitate focal contacts with adjacent cells (48).

Profilin 1 and actin translocation at the active membrane, along with the formation of podosome-mediated intercellular contacts and exosomal secretion, likely affect the high efficiency of HIV-1 infection that occurs with cell-cell fusion. In addition, increased secretion of profilin 1 in exosomes preceding syncytium formation indicates a possible role of the protein as a regulator of focal adhesions, actin stress filament orientation, and facilitation of giant cell formation. Similar profilin 1 secretion dynamics and RT activity in the HIV-1-infected culture fluids further support this hypothesis. During infection of HEp-2 cells by respiratory syncytial virus, profilin 1 plays a central role in virus-mediated stress fiber formation, as well as virion maturation and cellular fusion (49).

Although occurrence of giant cells has been described in granulomas and other inflammatory reactions since the middle of last century (50), their role in disease remains undefined. Cellular fusion has been shown to occur in the course of chronic inflammation mediated by IL-4 and IL-13 through a macrophage mannose receptor pathway. This model proposes cellular fusion and MGC formation as a necessity for increased phagocytosis of glycoproteins and microorganisms bearing terminal mannose, fucose, or glucose residues (51). This idea is complementary to other proposed models that describe cellular fusion as a prerequisite for increased contact area with the pathogen at the phagosomal site (52). Indeed, giant cells have been recognized previously as an end product of overwhelmed phagocytosis favored by high temperatures and acidic pH (53) and vessels for physical containment of the pathogen (54). We propose that HIV-1 accelerates macrophage differentiation and giant cell formation at the peak of viral replication and is reflective of a vain attempt of the cell to affect maximal pathogen clearance. We demonstrated that MGC possess newly reorganized actin networks and reduced membrane recycling. Images reveal elevated recruitment of profilin 1 and actin from the plasma membrane to the perinuclear region and profilin 1 and F-actin heavy staining of the nuclei. Based on these data, it is likely that profilin 1 along with actin, recently shown to participate in gene transcription and mRNA splicing (55, 56), may represent another check point in expression of viral proteins. Actin and profilin 1 structural modifications leading to their nuclear localization will be thoroughly investigated in future studies.

Culture fluids of MDM treated with latrunculin A (G-actin sequestering drug) revealed similar viral load compared with the infected controls (data not shown). The efficacy of its action, as well as the simplicity and specificity of latrunculin A interaction with G-actin, has made it a compound of choice, supplanting the classic actin-depolymerizing drug cytochalasin-D. A subpool of cytoplasmic G-actin may also be sequestered by latrunculin A, which explains the disruption of actin recruitment to the cellular interface and delayed MGC formation. Others have reported similar effects of actin depolymerizing drugs in cells infected with equine infectious anemia virus (57). Sequestration of actin by latrunculin A may exert its effects at multiple stages of the HIV-1 life cycle as the actin network is closely involved in viral entry and transport of the reverse transcription complexes to the nucleus, as well as progeny virion assembly and release (58).

Profilin plays important roles in cell growth and function (59, 60, 61). Its ability to bind actin (62), P-L-P (28, 29), a plethora of other proteins related to the microfilament system (63, 64), and to polyphosphoinositides (65) suggests that profilin 1 is involved in signal transduction and may link transmembrane signaling to the control of the microfilament system. The P-L-P binding site creates a stearic hindrance for some of profilin 1-binding partners that are involved in signaling events leading to the translocation of profilin 1 to distinct cellular regions, including the plasma membrane (28, 29, 30). P-L-P became our inhibitor of choice due to its high binding specificity to profilin 1 and minimal toxicity to the MDM. Complete inhibition of profilin 1 expression by siRNA was not preferred due to the suppressive effects of profilin 1 interaction with >50 profilin 1-binding partners and signal transduction intermediates (30).

HIV-1 may control temporal actin and profilin interactions, as well as shuttling to distinct cellular compartments, through the modulation of multiple posttranslational modification tags on these proteins. Both profilin 1 and actin posses the structural characteristics to acquire phosphorylation (65, 66) and acetylation (67) tags depending on the signaling event. Viruses, such as vaccinia and human T cell leukemia virus, exploit the actin cytoskeleton for efficient cell-to-cell spread through elaborate orchestration of signaling events that initiate with the Src-mediated tyrosine phosphorylation of viral proteins and actin-binding partners, including profilin 1 and Arp2/3 complexes leading to actin polymerization (68, 69, 70, 71). Evidence for HIV-1 exploitation of a similar mechanism is emerging. Recent studies have shown HIV-1 alters microtubule organization to build a "virological synapse" promoting efficient cell-to-cell spread and rapid dissemination of the progeny virus within the secondary lymphoid tissues in vivo (72). Our data hints that HIV-1 may recruit the actin skeleton to establish intercellular contacts and facilitate cell-to-cell progeny virion dissemination.

In conclusion, a link between the HIV-1-infected macrophage cytoskeleton and cell fusion was established. We demonstrated that viral replication and MGC formation is dependent on profilin 1-mediated actin stress filaments and the podosome-mediated intercellular network. These findings raise opportunities for therapeutic strategies that serve to interrupt secretion of the cytoskeleton and in this way can affect pivotal mechanisms for viral dissemination.


   Acknowledgments
 
We thank Dr. Gloria Borgstahl (Eppley Institute, University of Nebraska Medical Center) for providing purified actin and profilin 1. We thank Drs. S. Gorantla, H. Dou, L. Poluektova, J. Taylor, and S. Kraft for their assistance with the confocal microscopy studiesI, and L. Wilkie for FACS analyses. Dr. R. L. Mosley and R. Taylor are acknowledged for the careful reading of the manuscript. The help of T. Bargar and Dr. J. Rodriguez-Sierra in electron microscopy is appreciated.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported, in part, by National Institutes of Health Grants 2 R37 NS36126, 1 P01 NS31492, 1 P01 NS043985-01, 5 P01 MH64570-03, P20 RR15635 (to H.E.G.), and R21 MH75489 (to P.C.). Back

2 Address correspondence and reprint requests to Dr. Howard E. Gendelman, Center for Neurovirology and Neurodegenerative Disorders, University of Nebraska Medical Center 985880, Nebraska Medical Center, Omaha, NE 68198. E-mail address: hegendel{at}unmc.edu Back

3 Abbreviations used in this paper: MP, mononuclear phagocyte; GCI, Giant Cell Index; LC-MS/MS, liquid chromatography mass spectrometry; MDM, monocyte-derived macrophage; MGC, multinucleated giant cell; MOI, multiplicity of infection; P-L-P, poly-L-proline; RT, reverse transcriptase; SELDI-TOF, surface-enhanced laser desorption ionization time-of-flight; WCX2, weak cation exchange. Back

Received for publication December 9, 2006. Accepted for publication March 13, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Meuret, G., G. Hoffmann. 1973. Monocyte kinetic studies in normal and disease states. Br. J. Haematol. 24: 275-285. [Medline]
  2. van Furth, R., W. Sluiter. 1986. Distribution of blood monocytes between a marginating and a circulating pool. J. Exp. Med. 163: 474-479. [Abstract/Free Full Text]
  3. Devadas, K., N. J. Hardegen, L. M. Wahl, I. K. Hewlett, K. A. Clouse, K. M. Yamada, S. Dhawan. 2004. Mechanisms for macrophage-mediated HIV-1 induction. J. Immunol. 173: 6735-6744. [Abstract/Free Full Text]
  4. Peng, G., K. J. Lei, W. Jin, T. Greenwell-Wild, S. M. Wahl. 2006. Induction of APOBEC3 family proteins, a defensive maneuver underlying interferon-induced anti-HIV-1 activity. J. Exp. Med. 203: 41-46. [Abstract/Free Full Text]
  5. Ischiropoulos, H., L. Zhu, J. S. Beckman. 1992. Peroxynitrite formation from macrophage-derived nitric oxide. Arch. Biochem. Biophys. 298: 446-451. [Medline]
  6. Epstein, L. G., L. R. Sharer, J. Goudsmit. 1988. Neurological and neuropathological features of human immunodeficiency virus infection in children. Ann. Neurol. 23: (Suppl.):S19-S23. [Medline]
  7. Freed, E. O.. 2004. Mechanisms of enveloped virus release. Virus Res. 106: 85-86. [Medline]
  8. Peters, P. J., J. Bhattacharya, S. Hibbitts, M. T. Dittmar, G. Simmons, J. Bell, P. Simmonds, P. R. Clapham. 2004. Biological analysis of human immunodeficiency virus type 1 R5 envelopes amplified from brain and lymph node tissues of AIDS patients with neuropathology reveals two distinct tropism phenotypes and identifies envelopes in the brain that confer an enhanced tropism and fusigenicity for macrophages. J. Virol. 78: 6915-6926. [Abstract/Free Full Text]
  9. Iyengar, S., J. E. Hildreth, D. H. Schwartz. 1998. Actin-dependent receptor colocalization required for human immunodeficiency virus entry into host cells. J. Virol. 72: 5251-5255. [Abstract/Free Full Text]
  10. Collman, R. G.. 2003. HIV-1 Env-chemokine receptor interactions in primary human macrophages: entry and beyond. Res. Initiat. Treat. Action 8: 6-9. [Medline]
  11. Kedzierska, K., S. M. Crowe, S. Turville, A. L. Cunningham. 2003. The influence of cytokines, chemokines and their receptors on HIV-1 replication in monocytes and macrophages. Rev. Med. Virol. 13: 39-56. [Medline]
  12. Goldberg, M. B.. 2001. Actin-based motility of intracellular microbial pathogens. Microbiol. Mol. Biol. Rev. 65: 595-626. [Abstract/Free Full Text]
  13. Rogan, M. P., P. Geraghty, C. M. Greene, S. J. O’Neill, C. C. Taggart, N. G. McElvaney. 2006. Antimicrobial proteins and polypeptides in pulmonary innate defence. Respir. Res. 7: 29[Medline]
  14. Fackler, O. T., H. G. Krausslich. 2006. Interactions of human retroviruses with the host cell cytoskeleton. Curr. Opin. Microbiol. 9: 409-415. [Medline]
  15. Jolly, C., K. Kashefi, M. Hollinshead, Q. J. Sattentau. 2004. HIV-1 cell to cell transfer across an Env-induced, actin-dependent synapse. J. Exp. Med. 199: 283-293. [Abstract/Free Full Text]
  16. Gendelman, H. E., J. M. Orenstein, M. A. Martin, C. Ferrua, R. Mitra, T. Phipps, L. A. Wahl, H. C. Lane, A. S. Fauci, D. S. Burke, et al 1988. Efficient isolation and propagation of human immunodeficiency virus on recombinant colony-stimulating factor 1-treated monocytes. J. Exp. Med. 167: 1428-1441. [Abstract/Free Full Text]
  17. Kalter, D. C., M. Nakamura, J. A. Turpin, L. M. Baca, D. L. Hoover, C. Dieffenbach, P. Ralph, H. E. Gendelman, M. S. Meltzer. 1991. Enhanced HIV replication in macrophage colony-stimulating factor-treated monocytes. J. Immunol. 146: 298-306. [Abstract]
  18. Mosmann, T.. 1983. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. Immunol. Methods 65: 55-63. [Medline]
  19. Mor-Vaknin, N., A. Punturieri, K. Sitwala, D. M. Markovitz. 2003. Vimentin is secreted by activated macrophages. Nat. Cell Biol. 5: 59-63. [Medline]
  20. Enose, Y., C. J. Destache, A. L. Mack, J. R. Anderson, F. Ullrich, P. S. Ciborowski, H. E. Gendelman. 2005. Proteomic fingerprints distinguish microglia, bone marrow, and spleen macrophage populations. Glia 51: 161-172. [Medline]
  21. Yu, L. R., T. P. Conrads, T. Uo, Y. Kinoshita, R. S. Morrison, D. A. Lucas, K. C. Chan, J. Blonder, H. J. Issaq, T. D. Veenstra. 2004. Global analysis of the cortical neuron proteome. Mol. Cell. Proteomics 3: 896-907. [Abstract/Free Full Text]
  22. Ciborowski, P., I. Kadiu, W. Rozek, L. Smith, K. Bernhardt, M. Fladseth, M. Ricardo-Dukelow, and H. E. Gendelman. Investigating the human immunodeficiency virus type 1-infected monocyte-derived macrophage secretome. Virology. In press.
  23. Chertova, E., O. Chertov, L. V. Coren, J. D. Roser, C. M. Trubey, J. W. Bess, Jr, R. C. Sowder, II, E. Barsov, B. L. Hood, R. J. Fisher, et al 2006. Proteomic and biochemical analysis of purified human immunodeficiency virus type 1 produced from infected monocyte-derived macrophages. J. Virol. 80: 9039-9052. [Abstract/Free Full Text]
  24. Misumi, S., T. Fuchigami, N. Takamune, I. Takahashi, M. Takama, S. Shoji. 2002. Three isoforms of cyclophilin A associated with human immunodeficiency virus type 1 were found by proteomics by using two-dimensional gel electrophoresis and matrix-assisted laser desorption ionization-time of flight mass spectrometry. J. Virol. 76: 10000-10008. [Abstract/Free Full Text]
  25. Ott, D. E., L. V. Coren, B. P. Kane, L. K. Busch, D. G. Johnson, R. C. Sowder, II, E. N. Chertova, L. O. Arthur, L. E. Henderson. 1996. Cytoskeletal proteins inside human immunodeficiency virus type 1 virions. J. Virol. 70: 7734-7743. [Abstract]
  26. Thery, C., M. Boussac, P. Veron, P. Ricciardi-Castagnoli, G. Raposo, J. Garin, S. Amigorena. 2001. Proteomic analysis of dendritic cell-derived exosomes: a secreted subcellular compartment distinct from apoptotic vesicles. J. Immunol. 166: 7309-7318. [Abstract/Free Full Text]
  27. Sotiropoulos, A., D. Gineitis, J. Copeland, R. Treisman. 1999. Signal-regulated activation of serum response factor is mediated by changes in actin dynamics. Cell 98: 159-169. [Medline]
  28. Mahoney, N. M., D. A. Rozwarski, E. Fedorov, A. A. Fedorov, S. C. Almo. 1999. Profilin binds proline-rich ligands in two distinct amide backbone orientations. Nat. Struct. Biol. 6: 666-671. [Medline]
  29. Lindberg, U., C. E. Schutt, E. Hellsten, A. C. Tjader, T. Hult. 1988. The use of poly (L-proline)-Sepharose in the isolation of profilin and profilactin complexes. Biochim. Biophys. Acta 967: 391-400. [Medline]
  30. Witke, W.. 2004. The role of profilin complexes in cell motility and other cellular processes. Trends Cell Biol. 14: 461-469. [Medline]
  31. Pontow, S. E., N. V. Heyden, S. Wei, L. Ratner. 2004. Actin cytoskeletal reorganizations and coreceptor-mediated activation of rac during human immunodeficiency virus-induced cell fusion. J. Virol. 78: 7138-7147. [Abstract/Free Full Text]
  32. Miki, H., T. Sasaki, Y. Takai, T. Takenawa. 1998. Induction of filopodium formation by a WASP-related actin-depolymerizing protein N-WASP. Nature 391: 93-96. [Medline]
  33. Camera, P., J. S. da Silva, G. Griffiths, M. G. Giuffrida, L. Ferrara, V. Schubert, S. Imarisio, L. Silengo, C. G. Dotti, F. Di Cunto. 2003. Citron-N is a neuronal Rho-associated protein involved in Golgi organization through actin cytoskeleton regulation. Nat. Cell Biol. 5: 1071-1078. [Medline]
  34. Parast, M. M., C. A. Otey. 2000. Characterization of palladin, a novel protein localized to stress fibers and cell adhesions. J. Cell Biol. 150: 643-656. [Abstract/Free Full Text]
  35. Reinhard, M., K. Giehl, K. Abel, C. Haffner, T. Jarchau, V. Hoppe, B. M. Jockusch, U. Walter. 1995. The proline-rich focal adhesion and microfilament protein VASP is a ligand for profilins. EMBO J. 14: 1583-1589. [Medline]
  36. Luzio, J. P., B. M. Mullock, P. R. Pryor, M. R. Lindsay, D. E. James, R. C. Piper. 2001. Relationship between endosomes and lysosomes. Biochem. Soc. Trans. 29: 476-480. [Medline]
  37. de Gassart, A., C. Geminard, B. Fevrier, G. Raposo, M. Vidal. 2003. Lipid raft-associated protein sorting in exosomes. Blood 102: 4336-4344. [Abstract/Free Full Text]
  38. Witke, W., A. V. Podtelejnikov, A. Di Nardo, J. D. Sutherland, C. B. Gurniak, C. Dotti, M. Mann. 1998. In mouse brain profilin I and profilin II associate with regulators of the endocytic pathway and actin assembly. EMBO J. 17: 967-976. [Medline]
  39. Dong, J., B. Radau, A. Otto, E. Muller, C. Lindschau, P. Westermann. 2000. Profilin I attached to the Golgi is required for the formation of constitutive transport vesicles at the trans-Golgi network. Biochim. Biophys. Acta 1497: 253-260. [Medline]
  40. Kocks, C.. 1994. Intracellular motility: profilin puts pathogens on the actin drive. Curr. Biol. 4: 465-468. [Medline]
  41. Theriot, J. A., J. Rosenblatt, D. A. Portnoy, P. J. Goldschmidt-Clermont, T. J. Mitchison. 1994. Involvement of profilin in the actin-based motility of L. monocytogenes in cells and in cell-free extracts. Cell 76: 505-517. [Medline]
  42. Egile, C., T. P. Loisel, V. Laurent, R. Li, D. Pantaloni, P. J. Sansonetti, M. F. Carlier. 1999. Activation of the CDC42 effector N-WASP by the Shigella flexneri IcsA protein promotes actin nucleation by Arp2/3 complex and bacterial actin-based motility. J. Cell Biol. 146: 1319-1332. [Abstract/Free Full Text]
  43. Jay, D. G.. 2000. The clutch hypothesis revisited: ascribing the roles of actin-associated proteins in filopodial protrusion in the nerve growth cone. J. Neurobiol. 44: 114-125. [Medline]
  44. Gaidarov, I., F. Santini, R. A. Warren, J. H. Keen. 1999. Spatial control of coated-pit dynamics in living cells. Nat. Cell Biol. 1: 1-7. [Medline]
  45. Pelkmans, L., D. Puntener, A. Helenius. 2002. Local actin polymerization and dynamin recruitment in SV40-induced internalization of caveolae. Science 296: 535-539. [Abstract/Free Full Text]
  46. Lehmann, M. J., N. M. Sherer, C. B. Marks, M. Pypaert, W. Mothes. 2005. Actin- and myosin-driven movement of viruses along filopodia precedes their entry into cells. J. Cell Biol. 170: 317-325. [Abstract/Free Full Text]
  47. Lidke, D. S., K. A. Lidke, B. Rieger, T. M. Jovin, D. J. Arndt-Jovin. 2005. Reaching out for signals: filopodia sense EGF and respond by directed retrograde transport of activated receptors. J. Cell Biol. 170: 619-626. [Abstract/Free Full Text]
  48. Seabra, M. C., E. Coudrier. 2004. Rab GTPases and myosin motors in organelle motility. Traffic 5: 393-399. [Medline]
  49. Bitko, V., A. Oldenburg, N. E. Garmon, S. Barik. 2003. Profilin is required for viral morphogenesis, syncytium formation, and cell-specific stress fiber induction by respiratory syncytial virus. BMC Microbiol. 3: 9[Medline]
  50. Langhans, T.. 1868. Ueber Riesenzellen mit wandstandigen Kernen in Tuberkeln und die fibrosee Form des Tuberkels. Virchows Arch. Pathol. Anat. 42: 382-404.
  51. DeFife, K. M., C. R. Jenney, E. Colton, J. M. Anderson. 1999. Cytoskeletal and adhesive structural polarizations accompany IL-13-induced human macrophage fusion. J. Histochem. Cytochem. 47: 65-74. [Abstract/Free Full Text]
  52. McNally, A. K., J. M. Anderson. 2005. Multinucleated giant cell formation exhibits features of phagocytosis with participation of the endoplasmic reticulum. Exp. Mol. Pathol. 79: 126-135. [Medline]
  53. Anderson, J. M., K. Defife, A. McNally, T. Collier, C. Jenney. 1999. Monocyte, macrophage and foreign body giant cell interactions with molecularly engineered surfaces. J. Mater. Sci. Mater. Med. 10: 579-588. [Medline]
  54. Most, J., L. Spotl, G. Mayr, A. Gasser, A. Sarti, M. P. Dierich. 1997. Formation of multinucleated giant cells in vitro is dependent on the stage of monocyte to macrophage maturation. Blood 89: 662-671. [Abstract/Free Full Text]
  55. Stuven, T., E. Hartmann, D. Gorlich. 2003. Exportin 6: a novel nuclear export receptor that is specific for profilin actin complexes. EMBO J. 22: 5928-5940. [Medline]
  56. Skare, P., J. P. Kreivi, A. Bergstrom, R. Karlsson. 2003. Profilin I colocalizes with speckles and Cajal bodies: a possible role in pre-mRNA splicing. Exp. Cell Res. 286: 12-21. [Medline]
  57. Chen, C., O. A. Weisz, D. B. Stolz, S. C. Watkins, R. C. Montelaro. 2004. Differential effects of actin cytoskeleton dynamics on equine infectious anemia virus particle production. J. Virol. 78: 882-891. [Abstract/Free Full Text]
  58. Bukrinskaya, A., B. Brichacek, A. Mann, M. Stevenson. 1998. Establishment of a functional human immunodeficiency virus type 1 (HIV-1) reverse transcription complex involves the cytoskeleton. J. Exp. Med. 188: 2113-2125. [Abstract/Free Full Text]
  59. Machesky, L. M., T. D. Poland. 1993. Profilin as a potential mediator of membrane-cytoskeleton communication. Trends Cell Biol. 3: 381-385. [Medline]
  60. Cooley, L., E. Verheyen, K. Ayers. 1992. Chickadee encodes a profilin required for intercellular cytoplasm transport during Drosophila oogenesis. Cell 69: 173-184. [Medline]
  61. Haarer, B. K., S. S. Brown. 1990. Structure and function of profilin. Cell Motil. Cytoskeleton 17: 71-74. [Medline]
  62. Carlsson, L., L. E. Nystrom, I. Sundkvist, F. Markey, U. Lindberg. 1977. Actin polymerizability is influenced by profilin, a low molecular weight protein in non-muscle cells. J. Mol. Biol. 115: 465-483. [Medline]
  63. Gertler, F. B., K. Niebuhr, M. Reinhard, J. Wehland, P. Soriano. 1996. Mena,