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1α,25-Dihydroxyvitamin D3-induced Monocyte Antimycobacterial Activity Is Regulated by Phosphatidylinositol 3-Kinase and Mediated by the NADPH-dependent Phagocyte Oxidase*

Open AccessPublished:September 21, 2001DOI:https://doi.org/10.1074/jbc.M102876200
      We investigated the basis for the induction of monocyte antimycobacterial activity by 1α,25-dihydroxyvitamin D3 (D3). As expected, incubation of Mycobacterium tuberculosis-infected THP-1 cells or human peripheral blood, monocyte-derived macrophages with hormone resulted in the induction of antimycobacterial activity. This effect was significantly abrogated by pretreatment of cells with either of the phosphatidylinositol 3-kinase (PI 3-K) inhibitors, wortmannin or LY294002, or with antisense oligonucleotides to the p110 subunit of PI 3-Kα. Cells infected with M. tuberculosisalone or incubated with D3 alone produced little or undetectable amounts of superoxide anion (O⨪2). In contrast, exposure of M. tuberculosis-infected cells to D3 led to significant production of O⨪2, and this response was eliminated by either wortmannin, LY294002, or p110 antisense oligonucleotides. As was observed for PI 3-K inactivation, the reactive oxygen intermediate scavenger, 4-hydroxy-TEMPO, and degradative enzymes, polyethylene glycol coupled to either superoxide dismutase or catalase, also abrogated D3-induced antimycobacterial activity. Superoxide production by THP-1 cells in response to D3 required prior infection with liveM. tuberculosis, since exposure of cells to either killed M. tuberculosis or latex beads did not prime for an oxidative burst in response to subsequent hormone treatment. Consistent with these findings, redistribution of the cytosolic oxidase components p47phox and p67phox to the membrane fraction was observed in cells incubated with liveM. tuberculosis and D3 but not in response to combined treatment with heat-killed M. tuberculosis followed by D3. Redistribution of p47phox and p67phox to the membrane fraction in response to live M. tuberculosis and D3 was also abrogated under conditions where PI 3-K was inactivated. Taken together, these results indicate that D3-induced, human monocyte antimycobacterial activity is regulated by PI 3-K and mediated by the NADPH-dependent phagocyte oxidase.
      D3
      1α,25-dihydroxyvitamin D3
      MDM
      monocyte-derived macrophage(s)
      PI 3-K
      phosphatidylinositol 3-kinase
      Wm
      wortmannin
      LY
      LY294002
      O⨪2
      superoxide anion
      PEG
      polyethylene glycol
      SOD
      superoxide dismutase
      cat
      catalase
      VDR
      vitamin D receptor
      PI
      l-α-phosphatidylinositol
      l-NMMA
      l-N-monomethylarginine
      LDH
      lactate dehydrogenase
      NO
      nitric oxide
      LAM
      lipoarabinomannan
      WCL
      whole cell lysates
      ECF
      concentrated extracellular filtrate
      HBSS
      Hanks' balanced saline solution
      CFU
      colony-forming units
      PMA
      phorbol 12-myristate 13-acetate
      TUNEL
      terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling
      4-hydroxy-TEMPO
      4-hydroxy-2,2,6,6-tetramethylpiperidine-1-oxyl
      TLR-2
      toll-like receptor 2
      As the leading cause of death from any single bacterial infection in the world, tuberculosis represents a global health problem of paramount importance. Current estimates are that one-third of the world's population is infected with the etiological agentMycobacterium tuberculosis (
      • Kaufmann S.H.
      • van Embden J.D.
      ) and that the incidence of new cases with active disease is anticipated to rise from the current 7 million per year to 10 million per year by 2015 (
      • Day M.
      ). These statistics highlight the importance of developing new, more effective anti-tuberculous drugs, an effort dependent on acquiring more insights into host-pathogen interactions that determine the outcome of infection.
      M. tuberculosis primarily infects mononuclear phagocytes, where it resides and multiplies within a host-derived phagosome (
      • Armstrong J.A.
      • Hart P.D.
      ). Its success as a pathogen is largely attributable to its ability to evade or resist the multiplicity of antimicrobial mechanisms available to this host cell. The macrophage is not only the primary site for M. tuberculosis growth but also ordinarily provides the primary line of host defense against invading pathogens in its role as an effector of innate immunity. Macrophages use varied strategies to kill and destroy invading organisms, including production of reactive nitrogen and oxygen intermediates, phagosome maturation and acidification, fusion with lysosomes, exposure to defensins and host cell apoptosis (
      • Hingley-Wilson S.M.
      • Sly L.M.
      • Reiner N.E.
      • McMaster W.R.
      ). Augmentation of any of these processes during macrophage activation may contribute to control of disease.
      Vitamin D3 is a steroid hormone that regulates several cellular and physiological responses. The classical mechanism of action of D31 involves genomic signaling where hormone binds to the vitamin D receptor (VDR), a ligand-dependent transcription factor. The D3·VDR complex then translocates to the nucleus, where it directly regulates transcription by binding to the vitamin D response element consensus sequence located upstream of D3-activated genes (
      • Malloy P.J.
      • Feldman D.
      ). Recently, nongenomic signaling in response to D3 (i.e. cellular responses brought about independent of de novo transcription from a classical vitamin D response element) has been recognized to regulate important cellular processes (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ,
      • Gniadecki R.
      ,
      • Janis A.E.
      • Kaufmann S.H.E.
      • Schwartz R.H.
      • Pardoll D.M.
      ,
      • Phillips W.
      • Hamilton J.A.
      ,
      • Duncan R.
      • McConkey E.H.
      ).
      Vitamin D3 is known to possess a variety of immunomodulatory properties including effects on both myeloid and lymphoid cells (
      • Manolagas S.C.
      • Hustmyer F.G.
      • Yu X.P.
      ). Among these are its ability to induce the differentiation of immature myeloid cells into more mature monocytes and macrophages (
      • Abe E.
      • Miyaura C.
      • Sakagami H.
      • Takeda M.
      • Konno K.
      • Yamazaki T.
      • Yoshiki S.
      • Suda T.
      ,
      • Tanaka H.
      • Abe E.
      • Miyaura C.
      • Kuribayashi T.
      • Konno K.
      • Nishii Y.
      • Suda T.
      ,
      • Kreutz M.
      • Andreesen R.
      ,
      • Schwende H.
      • Fitzke E.
      • Ambs P.
      • Dieter P.
      ,
      • Zhang D.-E.
      • Hetherington C.J.
      • Gonzalez D.A.
      • Chen H.-M.
      • Tenen D.G.
      ). Several lines of evidence indicate that D3 regulates host resistance to M. tuberculosis. D3 deficiency and vitamin D receptor polymorphisms have been linked to increased susceptibility toM. tuberculosis and Mycobacterium leprae (
      • Davies P.D.
      ,
      • Roy S.
      • Frodsham A.
      • Saha B.
      • Hazra S.K.
      • Mascie-Taylor C.G.
      • Hill A.V.
      ,
      • Wilkinson R.J.
      • Llewelyn M.
      • Toossi Z.
      • Patel P.
      • Pasvol G.
      • Lalvani A.
      • Wright D.
      • Latif M.
      • Davidson R.N.
      ). In this regard, D3 productionin vivo is promoted by exposure to ultraviolet light, and this may provide a link between exposure to sunlight and antimycobacterial resistance mechanisms (
      • Davies P.D.
      ,
      • Crowle A.J.
      • Ross E.J.
      • May M.H.
      ). In addition, D3 has been shown to activate mononuclear phagocyte antimycobacterial activity (
      • Crowle A.J.
      • Ross E.J.
      • May M.H.
      ,
      • Rook G.A.
      • Steele J.
      • Fraher L.
      • Barker S.
      • Karmali R.
      • O'Riordan J.
      • Stanford J.
      ). Until now, the molecular basis through which D3 regulates host resistance to M. tuberculosis has not been identified.
      Class I phosphatidylinositol 3-kinase (PI 3-K) is a lipid kinase that phosphorylates the 3′-position of the inositol ring of phosphatidylinositol (PI) and its derivatives. It is composed of an 85-kDa Src homology 2 domain containing regulatory subunit (p85) and a 110-kDa catalytic subunit (p110). It is a multifunctional signaling molecule that has been implicated in a wide range of cellular processes including nuclear signaling, vesicle transport, organization of the cytoskeleton, cell growth, transformation, and survival (
      • Fry M.J.
      ,
      • Toker A.
      • Cantley L.G.
      ,
      • Fukui Y.
      • Ihara S.
      • Nagata S.
      ). It was recently found that PI 3-K is required for D3-induced cell differentiation in the human macrophage cell lines THP-1 and U937 and in peripheral blood monocytes (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ). In the latter study, vitamin D3 was observed to activate PI 3-K, and D3-induced expression of the monocyte differentiation markers, CD14 and CD11b, required PI 3-K. Furthermore, D3treatment induced the formation of a vitamin D receptor·PI 3-K complex, representing a novel nongenomic signaling pathway activated by D3. In light of these results, this study examined whether PI 3-K is involved in regulating D3-induced antimycobacterial activity and investigated the mechanistic basis for this effector function. The results obtained show that D3-induced monocyte resistance to M. tuberculosis is regulated by PI 3-K and that this effect is due to activation of the phagocyte NADPH oxidase.

      EXPERIMENTAL PROCEDURES

      Reagents and Chemicals

      RPMI 1640 and HBSS were from Stem Cell Technologies (Vancouver, Canada). Wortmannin, LY294002,l-NMMA, 1α,25-dihydroxyvitamin D3, 1α-hydroxyvitamin D3, and 25-hydroxyvitamin D3 were from Calbiochem.l-α-Phosphatidylinositol, phenylmethylsulfonyl fluoride, leupeptin, pepstatin A, aprotonin, microcystin, vanadate, ferricytochrome c, superoxide dismutase, latex beads, 4-hydroxy-TEMPO, PEG-catalase, and PEG-superoxide dismutase were from Sigma. Protein A-agarose, Griess reagent, and electrophoresis reagents were from Bio-Rad. [γ-32P]ATP was from Amersham Pharmacia Biotech. Phosphorothioate-modified oligonucleotides were synthesized by Life Technologies, Inc. Mannose-capped lipoarabinomannan was provided by D. John T. Belisle.

      Antibodies

      Mouse monoclonal anti-p85 UB93–3 was from Upstate Biotechnology Inc. (Lake Placid, NY). Mouse monoclonal anti-p110α was from Transduction Laboratories. Anti-p47phoxand anti-p67phox antibodies were prepared as described previously (
      • Allen L.A.
      • DeLeo F.R.
      • Gallois A.
      • Toyoshima S.
      • Suzuki K.
      • Nauseef W.M.
      ).

      Cell Lines

      The human promonocytic cell line THP-1 and murine macrophage-like cell line RAW264.7 were from the American Type Culture Collection. Cell lines were cultured in RPMI 1640 supplemented with 10% fetal calf serum (Hyclone), 2 mml-glutamine and maintained between 2 and 10 × 105 cells/ml. The human myeloid cell line THP-1 was used as a model for M. tuberculosis infection studies because of its similarity to human MDM (
      • Stokes R.W.
      • Doxsee D.
      ) in M. tuberculosis infection models and its availability for use.

      Isolation and Culture of Human Monocyte-derived Macrophages (MDM)

      Peripheral blood mononuclear cells were isolated as described previously (
      • Liu M.K.
      • Herrera-Velit P.
      • Brownsey R.W.
      • Reiner N.E.
      ). Mononuclear cells were allowed to adhere for 45 min at 37 °C in a humidified atmosphere with 5% CO2. Nonadherent cells were removed with three washes with HBSS. Adherent cells were maintained for 3 days at 37 °C in a humidified atmosphere with 5% CO2 prior to use for infections or treatments.

      Infections

      A virulent strain of M. tuberculosis (H37Rv) was grown to late log phase in Middlebrook 7H9 with OADC (Difco). Aliquots were frozen at −70 °C, and representative samples were thawed and evaluated for colony-forming units (CFUs) on Middlebrook 7H10 agar with OADC (Difco Laboratories). Heat-killed bacteria were prepared by heating at 80 °C for 2 h. Formaldehyde-fixed bacteria were prepared by fixation in 30% formaldehyde in methanol for 30 min. UV-irradiated bacteria were prepared by UV irradiation for 16 h. Each treatment reduced the viability of M. tuberculosis by greater than 5 logs. Cells were infected as described previously (
      • Hmama Z.
      • Gabathuler R.
      • Jefferies W.A.
      • de Jong G.
      • Reiner N.E.
      ). Briefly, prior to infection, THP-1 cells were seeded at a density of 105/cm2 in either six-well flat bottom or 10-cm diameter tissue culture plates (Becton Dickinson, Franklin Lakes, NJ) and allowed to adhere and differentiate in the presence of 20 ng/ml PMA at 37 °C in a 5% CO2 humidified atmosphere for 24 h. Cells were washed, and medium without PMA was replenished 4 h prior to the addition of bacteria. Prior to infection, bacteria were opsonized with fresh serum as described previously (
      • Hmama Z.
      • Gabathuler R.
      • Jefferies W.A.
      • de Jong G.
      • Reiner N.E.
      ). Monolayers were infected with opsonized M. tuberculosis at a 50:1 ratio. Latex beads were opsonized as for M. tuberculosis and incubated with cells at a ratio of 20:1. After 4 h, noningested bacilli or beads were removed by washing three times with HBSS, and medium was replenished. This resulted in infection of 80–90% of cells with 1–5 bacteria/cell. Infection was evaluated by Kinyoun staining (BBL Microbiology Systems, Cockeysville, MD). For latex beads, ∼90% of cells contained beads with 1–7 beads per cell.
      Immediately after infection, macrophages were treated with inhibitors at final concentrations as follows: LY, 14 μm; Wm, 50 nm, and l-NMMA, 8 μm. Cells were incubated with these inhibitors for 15 min at 37 °C, 5% CO2 prior to the addition of D3. 4-Hydroxy-TEMPO (0.1 mm), PEG-SOD (100 units/ml), or PEG-cat (100 units/ml) were added during the 4-h infection, and reagents remaining in the culture medium were washed away from treated cells along with noningested bacteria. Heat-inactivated PEG-SOD and PEG-cat were used as negative controls for the active enzymes. Vitamin D3 (1 μm) was added to monolayers after treatment with inhibitors and was left in the medium for the remainder of the experiment.

      In Vitro PI 3-K Assays

      In vitro PI 3-K assays were performed as described previously (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ).

      Determination of CFUs

      Colony-forming units were determined as described previously (
      • Crowle A.J.
      • Ross E.J.
      • May M.H.
      ). Bacilli were plated immediately after infection (time 0) and at 2 and 4 or 4 and 7 days after infection. Organisms were released in cold phosphate-buffered saline, 0.1% Triton X-100, serially diluted in Middlebrook 7H9, and 20 μl of three dilutions were plated in triplicate on Middlebrook 7H10 agar. CFUs were counted after 14 days of incubation and maintained for 21 days to ensure that no additional CFUs became visible.

      Sense and Antisense Phosphorothioate-modified Oligonucleotides

      Phosphorothioate-modified oligonucleotides were prepared and incorporated into cells as described previously (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ,
      • Volinia S.
      • Hiles I.
      • Ormondroyd E.
      • Nizetic D.
      • Antonacci R.
      • Rocchi M.
      • Waterfield M.D.
      ). Briefly, phosphorothioate-modified oligonucleotides to the α-isoform of the p110 subunit were synthesized and high pressure liquid chromatography-purified by Life Technologies, Inc. Oligonucleotides were phosphorothioate-modified to prevent intracellular degradation and were purified to remove incomplete synthesis products. 21-mers were produced to the human α isoform of the p110 subunit of PI 3-K, including the presumed translation initiation site in both sense and antisense directions with the following sequences: sense (5′-ATG CCT CCA AGA CCA TCA TCA-3′) and antisense (5′-TGA TGA TGG TCT TGG AGG CAT-3′). THP-1 cells (5 × 106) were resuspended in 500 μl of RPMI containing 2.5% LipofectAMINE (Life Technologies) and 5 μm phosphorothioate-modified oligonucleotides and incubated on a rotary shaker for 4 h at 37 °C prior to adherence and differentiation.
      NO3 Assays—Nitrite secreted by cells was measured using the Griess reagent according to the manufacturer's protocol (Sigma).

      O⨪2 Assays

      Superoxide assays were performed in triplicate as described previously (
      • DeLeo F.R.
      • Jutila M.A.
      • Quinn M.T.
      ) by measuring the superoxide dismutase inhibitable reduction of ferricytochrome c.

      Membrane and Cytosol Purification

      To separate membrane and cytosol fractions, 107 cells were resuspended in 1 ml of cold 5 mm HEPES, pH 7.4, and placed on ice for 20 min. Cell suspensions were homogenized on ice by 20 passages through a 22-gauge needle. Nuclei and intact cells were removed by centrifugation at 800 rpm for 2 min in a Microfuge. Sodium chloride was added to the supernatant to a final concentration of 0.15 m. Lysates were centrifuged at 15,000 rpm for 30 min. The resulting supernatant contained cytosolic proteins. The pellet was solubilized in 5 mm HEPES, pH 7.4, 1% Triton X-100, 0.15 mNaCl, 10% glycerol and incubated with rotation at 4 °C for 1 h. The resulting suspension was filter-sterilized through a 0.22-μm filter and contained membrane and membrane-associated proteins. 5′-nucleotidase and lactate dehydrogenase (LDH) assays were done to monitor for membrane contamination of cytosol and cytosolic contamination of membrane fractions, respectively.

      5′-Nucleotidase Assay

      5′-Nucleotidase activity was measured by the cleavage of phosphate from 5′-AMP (1.25 mm) in reaction buffer containing 10 mm HEPES, pH 7.4, 0.15m NaCl, 2 mm KCl, 2 mmMgCl2 for 15 min at room temperature. Liberated phosphate was measured by adding malachite green in 0.01% Tween 20 and allowing color to develop for 15 min at room temperature. The amount of phosphate released was determined from a standard curve.

      LDH Assays

      LDH assays were performed using an LDH assay kit from Sigma.

      SDS-Polyacrylamide Gel Electrophoresis and Western Immunoblotting

      SDS-polyacrylamide gel electrophoresis was performed by the method of Laemmli et al. (
      • Laemmli U.K.
      ). Membranes were developed by ECL as described previously (
      • Nandan D.
      • Reiner N.E.
      ).

      Apoptosis Assays

      Apoptosis was evaluated using the TUNEL assay (Calbiochem) including controls provided with the kit and incubation with actinomycin D as a positive control.

      Data Presentation and Statistical Analysis

      Data in graphs are expressed as means ± S.E. Statistical analyses for superoxide and nitrite assays were done by an unpaired Student's ttest. Comparisons for CFUs were done by analysis of variance for each time point. Differences were considered significant at a level ofp < 0.05.

      RESULTS

      D3-induced Antimycobacterial Activity in THP-1 Cells and Human Peripheral Blood MDM Is PI 3-Kinase-dependent

      Incubation of M. tuberculosis-infected THP-1 cells with the bioactive metabolite of vitamin D3 resulted in the induction of antimycobacterial activity. In contrast, no effect was observed using either the D3 precursor, 1-hydroxyvitamin D3 (1-OH D3) or an inactive analog, 25-hydroxyvitamin D3 (25-OH D3) (Fig.1 A). Whereas D3had no effect on the growth of M. tuberculosis in broth culture (data not shown), the number of CFUs recovered from THP-1 cells 4 and 7 days after treatment with D3 was 36 ± 3 and 20 ± 4%, respectively, compared with untreated, control cells (Fig. 1 A).
      Figure thumbnail gr1
      Figure 1Vitamin D3 induces antimycobacterial activity and PI 3-K activity in THP-1 cells. Panel A, THP-1 cells (1 × 106) were infected with a 50:1 ratio of opsonized M. tuberculosis strain H37Rv. After 4 h of co-incubation, cells were washed three times with HBSS, and the culture medium was replaced. THP-1 cells were left untreated (C) or were treated with 1 μm D3, 1-OH vitamin D3 (1-OH D3), or 25-OH vitamin D3(25-OH D3) immediately after infection. Numbersin parentheses above each barrepresent the percentage of CFUs observed for each treatment compared with control cells. THP-1 cells were lysed with cold phosphate-buffered saline containing 0.1% Triton X-100 immediately after infection and at 4 and 7 days after infection, and M. tuberculosiswas serially diluted in Middlebrook 7H9 medium and plated on Middlebrook 7H10 agar. CFUs were counted after 2 or 3 weeks of growth at 37 °C. CFUs were the same at time 0 for all treatment groups. Values represent the means ± S.E. of three independent experiments plated in triplicate. A, p < 0.01, comparing D3-treated versus control; N.S., not significantly different comparing treated versus control. Comparisons were made at each time point. Panel B, THP-1 cells (3 × 106) were left untreated (lane 1); differentiated with PMA (lane 2); co-incubated with 50:1 opsonized M. tuberculosisfor 2 (lane 3) or 4 h (lane 4); or infected with M. tuberculosisand treated with vitamin D3 for 10, 20, or 30 min (lanes 5, 7, and 9, respectively); or differentiated cells were treated with D3only for 10, 20, or 30 min (lanes 6,8, and 10, respectively). After treatment, cells were lysed and assayed in vitro for PI 3-K activity. The TLC shown is from one experiment representative of two independent experiments with similar results.
      Vitamin D3 has previously been shown to initiate a signaling pathway in human mononuclear phagocytes involving activation of PI 3-K (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ). Similarly, D3 treatment of M. tuberculosis-infected THP-1 cells activated PI 3-K (Fig.1 B, lanes 5, 7, and9 versus lane 4). To examine the role of PI 3-K in D3-induced antimycobacterial activity, the growth of M. tuberculosis was evaluated in D3-treated cells in the presence and absence of the PI 3-K inhibitors, LY and Wm, as well as in cells treated with antisense phosphorothioate-modified oligonucleotides to the p110 subunit of PI 3-Kα. Treatment with either LY or Wm inhibited PI 3-K activity in M. tuberculosis-infected, D3-treated THP-1 cells (Fig.2 A, lanes 4 and 6 versus lane 2). Treatment with either LY or Wm also significantly reduced the antimycobacterial effect of D3 at both day 4 and day 7 in THP-1 cells and day 2 and day 4 in human peripheral blood MDM (Fig. 2, B and C). The effect of LY or Wm was independent of effects on viability of the THP-1 cells. LDH activity was measured in culture supernatants of M. tuberculosis-infected, D3-treated cells 24 h after D3 treatment. Cells were treated with LY, Wm, or diluent. LDH activity at 25 °C was 124.4 units in diluent-treated or LY-treated culture supernatants and 125.9 units in Wm-treated culture supernatants. Analysis of apoptosis in similarly treated cells by TUNEL assay revealed 19.1% TUNEL-positive cells in control samples, 18.5% in LY-treated cells, and 17.2% in Wm-treated cells.
      Figure thumbnail gr2
      Figure 2PI 3-kinase regulates the induction of antimycobacterial activity by vitamin D3. Panel A, in vitro PI 3-K activity was measured inM. tuberculosis-infected THP-1 cells (3 × 106). Infected cells were left untreated (lane 1), treated with D3 (lane 2), treated with PI 3-K inhibitor LY (14 μm) or Wm (50 nm; lanes 3 and5), or treated with LY or Wm followed by the addition of D3 (lanes 4 and 6).Panel B, PI 3-K inhibitor LY or Wm were added to THP-1 cells immediately prior to treatment with D3. Panel C, LY or Wm was added to human MDM immediately prior to treatment with D3. Vitamin D3-treated (1 μm) cells are indicated by a plus sign. CFUs were determined as described in Fig. . Values represent the means ± S.E. of two (C) or three (B) independent experiments plated in triplicate. Numbers inparentheses above each bar represent the percentage of CFUs observed for each treatment compared with control cells. A, p < 0.01, comparing D3-treated versus control cells; B,p < 0.01 comparing LY or Wm + D3 versus D3 alone. Neither LY + D3 nor Wm + D3 was significantly different from cells treated with either inhibitor alone; C, p < 0.04 comparing LY or Wm + D3 versus D3-treated cells. Neither LY + D3 nor Wm + D3 was significantly different from cells treated with either inhibitor alone; N.S., not significantly different comparing treatment versus control. Comparisons were made at each time point.
      Treatment with antisense, but not sense, oligonucleotides to the p110α subunit of PI 3-K eliminated the expression of this protein subunit in M. tuberculosis-infected, D3-treated THP-1 cells (Fig.3 A, lane A versus lane S). Further, treatment of M. tuberculosis-infected THP-1 cells with p110 antisense oligonucleotides also significantly attenuated the antimycobacterial action of D3. At day 4, M. tuberculosis growth was restored from 37 ± 3 to 78 ± 18% of control cells, and at day 7 growth was restored from 21 ± 2 to 77 ± 9% of control cells (Fig. 3 B). Treatment with antisense oligonucleotides to the p110 subunit of PI 3-Kα did not alter either the infection rate or M. tuberculosis growth in non-D3-treated THP-1 cells (Fig. 3 B). Furthermore, the reconstitution of CFUs by antisense to the p110α isoform was as effective as was the use of either nonspecific inhibitor, LY or Wm. Taken together, these findings suggest that the antimycobacterial action of D3 is regulated by class I PI 3-K, p85/p110α.
      Figure thumbnail gr3
      Figure 3Antisense mRNA to class I PI 3-K p110α reduces p110α expression and antimycobacterial activity in M. tuberculosis-infected, D3-treated THP-1 cells. Panel A, THP-1 cells were left untreated (C), treated with sense phosphorothioate-modified oligonucleotides to the predicted translation start site of PI 3-K p110α (S), or treated with antisense phosphorothioate-modified oligonucleotides to the predicted translation start site of PI 3-K p110α (A). THP-1 cells were then infected with M. tuberculosis and treated with D3 for 10 min. Whole cell lysates were separated by SDS-PAGE, Western blotted, and immunodetected with anti-p110α.Panel B, M. tuberculosis growth was evaluated in control cells or in cells pretreated with sense (S) or antisense (A) oligonucleotides to the PI 3-K p110α subunit. Pretreatment with phosphorothioate-modified oligonucleotides did not affect CFUs at time 0. Values represent the mean ± S.E. of four independent experiments plated in triplicate.Numbers in parentheses above eachbar represent the percentage of CFUs observed for each treatment compared with control cells. A, p < 0.01, comparing D3-treated versus control cells; B,p < 0.01 comparing sense versus antisense pretreatment in D3-treated cells; N.S., not significantly different comparing treatment versus control and in comparing treatment with sense oligonucleotides + D3 with D3-treated cells. Comparisons were made at each time point.

      D3-induced Antimycobacterial Activity in Human Macrophages Is Not Due to Nitric Oxide (NO) Production

      The potential role of NO in mediating the effects of D3 inM. tuberculosis-infected cells was examined using the inducible nitric oxide synthase inhibitor l-NMMA and by direct measurement of nitrite production. Treatment ofM. tuberculosis-infected THP-1 cells withl-NMMA did not reduce the antimycobacterial action of D3 (Fig. 4 A), and neither THP-1 cells nor human MDM produced nitrite in response to D3 (data not shown). In contrast, D3-induced antimycobacterial activity in M. tuberculosis-infected, murine RAW 274.1 cells was significantly reduced by pretreatment with l-NMMA. Consistent with the implication that NO contributed to D3-induced antimycobacterial activity in murine cells, D3 treatment of infected RAW cells resulted in significant secretion of nitrite (Fig. 4 B), and this was markedly reduced in the presence of l-NMMA (Fig. 4 B). Also, in contrast to human macrophages, PI 3-K inhibitors did not affect the induction of antimycobacterial activity in infected RAW cells (data not shown). These findings demonstrate that the antimycobacterial action of D3 is NO-independent in human macrophages, whereas the converse is true in murine cells. Moreover, in the latter, this process is independent of any effects of PI 3-K.
      Figure thumbnail gr4
      Figure 4Vitamin D3-induced antimycobacterial activity is not mediated by nitric oxide. Panel A, the nitric oxide inhibitor, l-NMMA, was added to THP-1 cells and RAW 264.7 cells prior to treatment with vitamin D3 and CFUs were determined as described in Fig. . D3 treatment is indicated by a plus sign; cells treated with l-NMMA are indicated byN. Values represent the means ± S.E. of three independent experiments plated in triplicate. Numbers inparentheses above each bar represent the percentage of CFUs observed for each treatment compared with control cells. A, p < 0.001, comparing treatmentversus control cells; B, p < 0.002 comparing treatment versus D3-treated cells; N.S., not significantly different comparing pretreatment withl-NMMA versus control and comparing treatment with l-NMMA + D3 with D3-treated THP-1 cells. Panel B, nitrite concentration in RAW 264.7 cells was measured at 4 and 7 days after treatment using Griess reagent. THP-1 cells did not secrete detectable amounts of nitrite under any condition (data not shown). Values represent the mean ± S.E. of three independent experiments performed in triplicate. A,p < 0.001, comparing treatment versuscontrol cells; B, p < 0.001 comparing treatmentversus D3-treated cells; C, p < 0.001 comparing pretreatment with l-NMMA versuscontrol cells; N.S., not significantly different comparing pretreatment with l-NMMA versus control cells. Comparisons were made at each time point.

      D3 Induces O⨪2 Production by Infected Human Macrophages in a PI 3-K-dependent Manner

      To examine further mechanisms accounting for the antimycobacterial action of D3, superoxide anion production by THP-1 cells and MDM was measured. M. tuberculosis-infected macrophages secreted large amounts of O⨪2 when treated with D3, and this effect was inhibited by LY and Wm (Fig.5, A and C) as well as by antisense phosphorothioate-modified oligonucleotides to the p110α subunit of PI 3-K (Fig. 5 B). Superoxide production occurred within the first 1 h following treatment with D3. There was no detectable superoxide produced by macrophages during the remainder of the experiment. M. tuberculosis infection alone induced the production of 20 ± 3 nmol of O⨪2/106 THP-1 cells/h, and similar values were observed for treatment with D3 alone. However, infection with M. tuberculosis followed by treatment with D3 was observed to result in the production of significantly more O⨪2 (88 nmol of O⨪2/106 macrophages/h). The amount of O⨪2 secreted by infected THP-1 cells in response to D3 was reduced to 41 ± 14 and 33 ± 13 nmol of O⨪2/106 macrophages/h by pretreatment with LY or Wm, respectively (Fig. 5 A). Antisense oligonucleotides to PI 3-K also reduced O⨪2 secretion by M. tuberculosis-infected, D3-treated THP-1 cells from 94 ± 3 to 17 ± 2 nmol/106 macrophages/h, while sense oligonucleotides had no effect (Fig. 5 B). Similar results were obtained using MDM (Fig. 5 C).M. tuberculosis infection alone induced the production of 28 ± 1 nmol of O⨪2/106macrophages/h, and D3 alone induced production of 16 ± 2 nmol of O⨪2/106 macrophages/h. However, infection followed by D3 treatment resulted in the production of significantly higher amounts of O⨪2(78 nmol of O⨪2/106 macrophages/h) by MDM. This amount was reduced to 36 ± 2 and 36 ± 3 nmol of O⨪2/106 macrophages/h by pretreatment with either LY or Wm, respectively. Taken together, these results suggest an interaction between M. tuberculosis infection and D3 in triggering the phagocyte oxidative burst in human MDM as well as in THP-1 cells and that this process is regulated by PI 3-K.
      Figure thumbnail gr5
      Figure 5D3 induces the phagocyte oxidative burst in M. tuberculosis-infected THP-1 cells in a PI 3-K-dependent manner. Superoxide secreted by macrophages in 1 h was determined by measuring the SOD-inhibitable reduction of ferricytochrome c. Panel A, superoxide secretion was evaluated in control THP-1 cells or THP-1 cells treated with PI 3-K inhibitor, LY (14 μm) or Wm (50 nm), immediately after infection with or without subsequent treatment with vitamin D3. Values represent the means ± S.E. of three independent experiments. Panel B, superoxide secretion was measured for control cells or cells pretreated with phosphorothioate-modified oligonucleotides to the predicted translational start site of the sense (S) or antisense (A) strand of the PI 3-K p110 subunit. Values represent the means ± S.E. from three independent experiments performed in triplicate. Panel C, superoxide secretion was evaluated in control MDM or cells pretreated with PI 3-K inhibitors, LY (14 μm) or Wm (50 nm). Values represent the means ± S.E. from two independent experiments performed in triplicate. A, p < 0.04, comparing treatmentversus M. tuberculosis-infected or D3-treated cells; B, p < 0.05 comparing treatment versus M. tuberculosis-infected, D3-treated cells; C,p < 0.001 comparing treatment versus M. tuberculosis-infected or D3-treated cells; D, p < 0.001 comparing pretreatment with antisense oligonucleotides versus sense oligonucleotides; E, p < 0.005 comparing treatmentversus M. tuberculosis-infected or D3-treated cells; F, p < 0.005 comparing treatment versus M. tuberculosisinfected, D3-treated cells; N.S., not significantly different comparing treatment versus M. tuberculosis infected or D3-treated cells.

      Phagocytosis per se Is Not Sufficient to Prime Cells for an Oxidative Burst in Response to D3

      Superoxide production was measured in cells treated with either live M. tuberculosis, latex beads, or killed M. tuberculosis alone or in combination with D3. As individual stimuli, live M. tuberculosis, latex beads, heat-killed M. tuberculosis, formaldehyde-fixed M. tuberculosis, UV-irradiatedM. tuberculosis, and D3 elicited comparable and only minimal amounts of O⨪2 production (less than 30 nmol of O⨪2/106 macrophages/h) by THP-1 cells (Fig. 6). However, infection with live M. tuberculosis primed cells for a marked increase in O⨪2 production in response to D3, and this was not observed with either killedM. tuberculosis or latex beads. These findings indicate that phagocytosis per se is not sufficient to prime cells and that viable M. tuberculosis actively contribute to the observed potentiation of the oxidative burst in response to D3.
      Figure thumbnail gr6
      Figure 6Phagocytosis is not sufficient to prime cells for an oxidative burst in response to D3. Superoxide production by THP-1 cells ingesting opsonized live M. tuberculosis strain H37Rv (TB), latex beads (LB), heat-killed (80 °C, 2 h) M. tuberculosis (HK), formaldehyde-fixed (30% formaldehyde in methanol, 30 min) M. tuberculosis(FF), or UV-irradiated (16 h) M. tuberculosis (UV), with or without the addition of D3 (1 μm) was examined. Superoxide secretion was determined by measuring the SOD-inhibitable reduction of ferricytochrome c in 1 h. The values shown are the means ± S.E. of two (for formaldehyde-fixed and UV-irradiated) or three independent experiments performed in triplicate. A,p < 0.01 comparing M. tuberculosis-infected, D3-treated THP-1 cells with cells infected with M. tuberculosis or treated with D3 alone; N.S., not significantly different comparing treatment with THP-1 cells infected with M. tuberculosis or treated with D3 alone.

      D3-induced O⨪2 Production Is Required for the Antimycobacterial Action of D3

      To assess the role of the phagocyte oxidative burst in the antimycobacterial action of D3, 4-hydroxy-TEMPO, PEG-SOD, and PEG-cat were used to examine whether killing of M. tuberculosiswas related to the production of reactive oxygen intermediates. Whereas none of these compounds alone affected the growth of M. tuberculosis in untreated cells, each of them significantly attenuated the ability of D3 to induce resistance toM. tuberculosis in THP-1 cells and MDM (Fig. 7, A and B). Heat-inactivated PEG-SOD or PEG-cat, however, had no effect on D3-induced anti-mycobacterial activity (data not shown). These results indicate that removal of reactive oxygen intermediates, either by scavenging (4-hydroxy-TEMPO) or enzymatically (PEG-cat or PEG-SOD) significantly nullifies the antimycobacterial action of D3.
      Figure thumbnail gr7
      Figure 7The oxygen radical scavenger, 4-hydroxy-TEMPO , and the hydrogen peroxide and superoxide degrading enzymes, cat and SOD , coupled to polyethylene glycol attenuate D3-induced antimycobacterial activity.4-Hydroxy-TEMPO, PEG-cat, or PEG-SOD was co-incubated with THP-1 cells (A) or MDM (B) during infection withM. tuberculosis. Growth of bacilli was evaluated as described in the legend to Fig. . Heat-inactivated PEG-cat or PEG-SOD did not affect D3-induced antimycobacterial activity. Vitamin D3-treated cells are indicated by aplus sign; cells treated with 4-hydroxy-TEMPO are indicated by a T; cells treated with PEG-catalase are indicated by a C; and cells treated with PEG-SOD are indicated by an S. Treatment with 4-hydroxy-TEMPO, PEG-cat, or PEG-SOD did not affect CFUs recovered at time 0. Values reported are the means ± S.E. of two (B) or three (A) independent experiments plated in triplicate. Numbers inparentheses above each bar represent the percentage of CFUs observed for each treatment compared with control cells. A, p < 0.001 comparing D3-treated with control cells; B, p < 0.03 comparing treatment to D3-treated cells. C,p < 0.02 comparing D3-treated with control cells; D, p < 0.04 comparing treatment with D3-treated cells; N.S., not significantly different from untreated control cells. Comparisons were made at each time point.

      D3 Induces the Association of Phagosome Oxidase Components with the Membrane Fraction in M. tuberculosis-infected THP-1 Cells

      Phagosome oxidase activity requires recruitment of the cytosolic components, p47phox and p67phox, to the membrane for assembly with flavocytochrome b 558(
      • Clark R.A.
      • Volpp B.D.
      • Leidal K.G.
      • Nauseef W.M.
      ,
      • DeLeo F.R.
      • Allen L.A.
      • Apicella M.
      • Nauseef W.M.
      ). The relative distribution of p47phox and p67phox in cytosolic and membrane fractions of stimulated cells was evaluated. Incubation of M. tuberculosis-infected cells with D3 resulted in a marked translocation of both p47phox and p67phox to the membrane fraction (Fig.8 A). Untreated and PMA-differentiated THP-1 cells had similar amounts of membrane-associated p47phox and p67phox. Incubation of cells with live M. tuberculosis alone for either 2 or 4 h resulted in minimal changes in the membrane association of p47phox and p67phox; however, subsequent treatment with D3 increased these amounts to 7.6- and 8.1-fold, respectively, over resting cells (Fig. 8 A, lane 5 versus lane 1). Exposure of cells to either heat-killed M. tuberculosisalone or followed by treatment with D3 resulted in minimal or no detectable increases in membrane levels of oxidase components (Fig. 8 A, lanes 6 and 7). Treatment of cells with either live or heat-killed M. tuberculosis alone or in combination with D3 did not bring about detectable changes in cytosolic levels of either p47phox or p67phox (Fig. 8 B), consistent with the relatively small fraction of p47phox and p67phox translocated during NADPH oxidase activation (
      • Clark R.A.
      • Volpp B.D.
      • Leidal K.G.
      • Nauseef W.M.
      ,
      • DeLeo F.R.
      • Allen L.A.
      • Apicella M.
      • Nauseef W.M.
      ).
      Figure thumbnail gr8
      Figure 8THP-1 cell cytosol and membrane immunoblots probed with anti-p47phox (upper panel) or anti-p67phox (lower panel). In panels A andB, membrane and cytosolic fractions, respectively, were purified from resting cells (lane 1), cells differentiated with PMA (lane 2), differentiated cells co-incubated with M. tuberculosis for 2 (lane 3) or 4 (lane 4) h, differentiated cells co-incubated with M. tuberculosis for 4 h followed by treatment with D3 (lane 5), differentiated cells co-incubated with heat-killed M. tuberculosis for 2 (lane 6) or 4 h (lane 7), differentiated cells co-incubated with heat-killedM. tuberculosis for 4 h followed by treatment with D3 (lane 8), and differentiated cells treated with D3 only (lane 9). In panels C and D, membrane fractions were purified from differentiated cells treated in the presence or absence of the PI 3-K inhibitor, LY or Wm, respectively. Shown are PMA-differentiated cells (lane 1), cells co-incubated with M. tuberculosis for 4 h (lane 2), D3-treated cells (lane 3), cells co-incubated with M. tuberculosis followed by D3 treatment (lane 4), PMA-differentiated cells treated with PI 3-K inhibitor (lane 5), cells treated with PI 3-K inhibitor followed by co-incubation with M. tuberculosis for 4 h (lane 6), cells treated with PI 3-K inhibitor followed by D3 treatment (lane 7), cells treated with PI 3-K inhibitor followed by co-incubation withM. tuberculosis for 4 h and then D3 treatment (lane 8), and cells co-incubated with M. tuberculosis followed by treatment with PI 3-K inhibitor and then D3(lane 9). Densitometry readings for each band were measured, and the values reported are averages from two independent experiments.

      D3-induced Translocation of Cytosolic Oxidase Components to the Membrane Fraction Is PI 3-K-dependent

      The redistribution of p47phoxand p67phox to the membrane fraction in response to infection with M. tuberculosis and treatment with D3 was markedly reduced in cells pretreated with either of the PI 3-K inhibitors, LY or Wm (Fig. 8, C andD). In M. tuberculosis-infected cells treated with D3, the amount of p47phox associated with the membrane increased 8.9-fold over control cells, and this was reduced to 2.4- and 3.2-fold, respectively, by treatment with LY either prior to infection or prior to D3 treatment (Fig.8 C, upper panel, lanes 8 and 9 versus lane 4). Similarly, levels of p67phox associated with the membrane increased 7.1-fold over PMA-differentiated cells, and this was reduced to 2.3- and 1.9-fold, respectively, by treatment with LY either prior to infection or prior to D3 treatment (Fig.8 C, lower panel, lanes 8 and 9 versus lane 4). The amount of p47phox associated with the membrane increased 6.3-fold over PMA-differentiated cells, and this was reduced to 1.2-fold by treatment with Wm either prior to infection or prior to D3 treatment (Fig. 8 D, upper panel, lanes 8 and 9 versus lane 4). The level of p67phox associated with the membrane increased 9.3-fold over PMA-differentiated cells, and this was reduced to 2.3- and 2.1-fold, respectively, by treatment with Wm either prior to infection or prior to D3 treatment (Fig. 8 D, lower panel, lanes 8 and 9 versus lane 4).

      Priming for the Phagocyte Oxidative Burst in Response to D3 by Purified M. tuberculosis LAM

      The ability of subcellular components of M. tuberculosis, in combination with D3, to bring about O⨪2 production by THP-1 cells was examined. Incubation of cells with 100 ng of purified M. tuberculosis LAM, WCL, or ECF elicited the production of 22 ± 3, 6 ± 2, or 1.2 ± 0.5 nmol of O⨪2/106 macrophages/h, respectively (Fig. 9). Whereas incubation with D3 alone elicited the production of only 17 ± 2 nmol of O⨪2/106 macrophages/h, pretreatment with LAM followed by D3 resulted in significantly enhanced production of O⨪2. In contrast, WCL primed minimally and ECF not at all for an enhanced oxidative burst in response to D3.
      Figure thumbnail gr9
      Figure 9Priming for the phagocyte oxidative burst in response to D3 by purified M. tuberculosis LAM. D3 induced superoxide production by THP-1 cells pretreated with LAM, WCL, or concentrated ECF. THP-1 cells (1 × 106) were treated with 100 ng of LAM, WCL, or ECF, and superoxide secretion by cells was examined with or without treatment with 1 μm vitamin D3. The values shown are the means ± S.E. of three independent experiments performed in triplicate. A, p < 0.01 comparing pretreatment with LAM or WCL followed by D3 treatment with D3-treated cells; N.S., not significantly different comparing pretreatment with ECF followed by D3 treatment with D3-treated cells.

      DISCUSSION

      The steroid hormone vitamin D3 has been linked to host resistance to tuberculosis since the mid-19th century (
      • Davies P.D.
      ), and the active metabolite, 1α,25-dihydroxyvitamin D3 has been shown to induce antimycobacterial activity in macrophages (
      • Crowle A.J.
      • Ross E.J.
      • May M.H.
      ,
      • Rook G.A.
      • Steele J.
      • Fraher L.
      • Barker S.
      • Karmali R.
      • O'Riordan J.
      • Stanford J.
      ). In addition, it has been found that the antimycobacterial efficacy of cholecalciferol metabolites correlated with their binding affinities to the VDR (
      • Rook G.A.
      • Steele J.
      • Fraher L.
      • Barker S.
      • Karmali R.
      • O'Riordan J.
      • Stanford J.
      ). Other evidence for the role of D3 in host resistance to tuberculosis has emerged recently. For example, D3 deficiency has been linked to increased susceptibility to tuberculosis, and evidence suggests that vitamin D receptor polymorphisms confer differential resistance to mycobacterial diseases including leprosy and tuberculosis (
      • Roy S.
      • Frodsham A.
      • Saha B.
      • Hazra S.K.
      • Mascie-Taylor C.G.
      • Hill A.V.
      ,
      • Wilkinson R.J.
      • Llewelyn M.
      • Toossi Z.
      • Patel P.
      • Pasvol G.
      • Lalvani A.
      • Wright D.
      • Latif M.
      • Davidson R.N.
      ,
      • Bellamy R.
      • Ruwende C.
      • Corrah T.
      • McAdam K.P.
      • Thursz M.
      • Whittle H.C.
      • Hill A.V.
      ). Whereas it is clear from these observations that D3 enhances host resistance to tuberculosis and other mycobacterial diseases, the underlying mechanism accounting for D3-mediated antimycobacterial activity in human cells and its regulation have been obscure.
      The findings of the present study indicate that monocyte antimycobacterial activity induced by 1α,25-dihydroxyvitamin D3 is mediated by the NADPH-dependent phagocyte oxidase and that this process is regulated by phosphatidylinositol 3-kinase. The conclusion that D3-induced antimycobacterial activity is due to activation of the phagocyte oxidative burst is based on three lines of evidence: (i) D3 treatment of M. tuberculosis-infected cells leads to a burst of O⨪2 production (Fig. 5, A and C); (ii) the reactive oxygen intermediate scavenger, 4-hydroxy-TEMPO, and enzymes that degrade reactive oxygen intermediates (PEG-cat and PEG-SOD) dramatically reduce the D3-mediated antimycobacterial effect in both THP-1 cells and MDM (Fig. 7,A and B); and (iii) the phagosome oxidase components, p47phox and p67phox, translocate to the membrane fraction in M. tuberculosis-infected cells upon treatment with D3, as would be expected during assembly of a functional oxidase (Fig. 8 A).
      Increased production of superoxide by infected cells in response to D3 was rapid and transient, whereas we were only able to detect a decrease in M. tuberculosis CFUs at either 2, 4, or 7 days following D3 treatment. This apparently delayed effect could reflect the production of low, undetectable levels of superoxide for longer periods of time. However, we believe a more likely possibility is that D3-induced antimycobacterial activity mediated by the NADPH oxidase acts synergistically by rendering M. tuberculosis more susceptible to other host factors that contribute to decreased CFUs observed over a longer time course. This model is consistent with the recent finding that NADPH oxidase-dependent killing of Salmonella in vivo was confined to the first few hours after phagocytosis. Reactive oxygen intermediates were essential to control infection but contributed at the very early stages of a multiphasic host response (
      • Vazquez-Torres A.
      • Jones-Carson J.
      • Mastroeni P.
      • Ischiropolous H.
      • Fang F.C.
      ).
      The conclusion that the oxidative burst is the principal mechanism by which D3 mediates antimycobacterial activity in human mononuclear phagocytes is consistent with genetic evidence linking the phagocyte oxidase to host resistance to mycobacteria. Thus, patients with chronic granulomatous disease, a heterogeneous group of genetic disorders involving functional inactivation of the phagocyte oxidase (
      • Babior B.M.
      ), show increased susceptibility to recurrent and life-threatening bacterial and fungal infections including infections due to mycobacteria (
      • Ohga S.
      • Ikeuchi K.
      • Kadoya R.
      • Okada K.
      • Miyazaki C.
      • Suita S.
      • Ueda K.
      ,
      • Allen D.M.
      • Chng H.H.
      ,
      • Chusid M.J.
      • Parrillo J.E.
      • Fauci A.S.
      ,
      • Gonzalez B.
      • Moreno S.
      • Burdach R.
      • Valenzuela M.T.
      • Henriquez A.
      • Ramos M.I.
      • Sorensen R.U.
      ).
      Vitamin D3 has been reported to stimulate directly human peripheral blood monocytes to generate an oxidative burst based upon the finding that it induced peripheral blood monocytes to secrete hydrogen peroxide (
      • Cohen M.S.
      • Mesler D.E.
      • Snipes R.G.
      • Gray T.K.
      ). In the present study, either D3 orM. tuberculosis acting alone triggered only modest oxidative burst activity. In respect to M. tuberculosis, this finding is consistent with previous reports showing only low levels of phagocyte oxidase activation in response to infection with mycobacteria (
      • Brett S.J.
      • Butler R.
      ,
      • Gordon A.H.
      • Hart P.D.
      ). In contrast to the results obtained with either agent acting alone, in the presence ofM. tuberculosis infection the oxidative burst in response to D3 was markedly augmented. These findings indicate that M. tuberculosis infection acted to prime cells to respond to D3, and this priming was specific for viable M. tuberculosis, since it was not observed in cells that had ingested either dead bacilli or latex beads.
      The precise mechanism for this priming effect by viable bacilli is unclear but probably involves the recruitment of one or more signaling pathways that act in conjunction with PI 3-K to prompt assembly of a functional oxidase complex. The finding that M. tuberculosis LAM was able to condition cells for an enhanced oxidative response to D3 (Fig. 9) provides a potential mechanism to account for the priming observed in response to infection with live M. tuberculosis. LAM is known to have pleiotropic effects including effects on multiple signaling pathways in macrophages (
      • Reiner N.E.
      ,
      • Knutson K.L.
      • Hmama Z.
      • Herrera-Velit P.
      • Rochford R.
      • Reiner N.E.
      ,
      • Maiti D.
      • Bhattacharyya A.
      • Basu J.
      ). The requirement for infection with live bacilli may reflect a need for de novo synthesis of LAM within cells, for the intracellular release and trafficking of LAM, or both. At the same time, although it was possible to show that LAM had the property of oxidase priming, these results do not establish that it is the actual in vivo priming factor or that it is the only priming factor elaborated by viable M. tuberculosis within cells.
      In activated murine macrophages, antimycobacterial activity has been shown to be due, at least in part, to the inducible synthesis of nitric oxide, and studies in inducible nitric oxide synthase knock out mice have suggested that this is an effector arm of the host response to M. tuberculosis (
      • Chan J.
      • Xing Y.
      • Magliozzo R.S.
      • Bloom B.R.
      ,
      • MacMicking J.D.
      • North R.J.
      • LaCourse R.
      • Mudgett J.S.
      • Shah S.K.
      • Nathan C.F.
      ). In the experiments reported above, detectable levels of nitrite were not produced by D3-treated, M. tuberculosis-infected THP-1 cells or MDM, and the NO inhibitor l-NMMA did not affect the induction of antimycobacterial activity by D3 in THP-1 cells (Fig. 4 A). This was in direct contrast to parallel observations made using the murine macrophage-like cell line RAW 274.1, which when infected with M. tuberculosis produced significant amounts of nitrite in response to D3 (Fig. 4 B) and in which the antimycobacterial effect of D3 was markedly attenuated by prior treatment with l-NMMA (Fig. 4 A). These results suggest that at least two independent mechanisms operate to induce antimycobacterial activity in D3-treated macrophages. In murine cells, inducible nitric oxide synthase operates independently of PI 3-K, whereas in human macrophages the NADPH-dependent phagocyte oxidative burst appears to be predominant and regulated by PI 3-K. Moreover, these findings are consistent with a recent report that examined the mechanisms involved in the antimycobacterial activity of toll-like receptor 2-activated macrophages (
      • Thoma-Uszynski S.
      • Stenger S.
      • Takeuchi O.
      • Ochoa M.T.
      • Engele M.
      • Sieling P.A.
      • Barnes P.F.
      • Rollinghoff M.
      • Bolcskei P.L.
      • Wagner M.
      • Akira S.
      • Norgard M.V.
      • Belisle J.T.
      • Godowski P.J.
      • Bloom B.R.
      • Modlin R.L.
      ). In these studies, TLR-2-activated murine macrophages demonstrated NO-dependent antimycobacterial activity, whereas TLR-2-activated antimycobacterial activity in human macrophages was NO-independent. No effector mechanism for TLR-2-activated antimycobacterial activity was defined in human cells (
      • Thoma-Uszynski S.
      • Stenger S.
      • Takeuchi O.
      • Ochoa M.T.
      • Engele M.
      • Sieling P.A.
      • Barnes P.F.
      • Rollinghoff M.
      • Bolcskei P.L.
      • Wagner M.
      • Akira S.
      • Norgard M.V.
      • Belisle J.T.
      • Godowski P.J.
      • Bloom B.R.
      • Modlin R.L.
      ). Our results presented here suggest that one possible effector mechanism is oxygen-dependent killing via activation of the phagocyte oxidase. This model is supported further by the findings that, when ligand-activated, the D3 receptor (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ) and TLR-2 (
      • Arbibe L.
      • Mira J.-P.
      • Teusch N.
      • Kline L.
      • Mausumee G.
      • Mackman N.
      • Godowski P.J.
      • Ulevitch R.J.
      • Knaus U.G.
      ) have in common the property of binding to PI 3-K. As we have shown, in M. tuberculosis-infected, D3-treated cells, this results in activation of the phagocyte oxidase. It seems highly likely that this would occur as well in TLR-2-activated cells infected with mycobacteria.
      Recently, there has been considerable interest in whether macrophage apoptosis is involved in the host response to mycobacteria and, if so, whether apoptosis could affect mycobacterial viability. Although results from various studies have been inconsistent (
      • Santucci M.B.
      • Amicosante M.
      • Cicconi R.
      • Montesano C.
      • Casarini M.
      • Giosuè S.
      • Bisetti A.
      • Colizzi V.
      • Fraziano M.
      ,
      • Kremer L.
      • Estaquier J.
      • Brandt E.
      • Ameisen J.C.
      • Locht C.
      ,
      • Placido R.
      • Mancino G.
      • Amendola A.
      • Mariani F.
      • Vendetti S.
      • Piacentini M.
      • Sanduzzi A.
      • Bocchino M.L.
      • Zembala M.
      • Colizzi V.
      ,
      • Durrbaum-Landmann I.
      • Gercken J.
      • Flad H.D.
      • Ernst M.
      ,
      • Keane J.
      • Remold H.G.
      • Kornfeld H.
      ,
      • Klingler K.
      • Tchou-Wong K.M.
      • Brandli O.
      • Aston C.
      • Kim R.
      • Chi C.
      • Rom W.N.
      ), we nevertheless considered the possibility that D3-induced antimycobacterial activity could be due to the induction of apoptosis. Rates of apoptosis in THP-1 cells incubated with D3 alone for either 1 or 4 days were at background levels (1–2%). In cells infected with M. tuberculosis alone, apoptosis was detected at frequencies of 13 and 15%, respectively, at 1 and 4 days postinfection, and these rates were not affected by the addition of D3. Similar results were found using human MDM. Treatment with D3 alone for 1 or 4 days resulted in levels of apoptosis similar to untreated cells (4 or 6%, respectively). Cells infected with M. tuberculosis alone showed apoptosis in 17 and 20% of cells at 1 and 4 days postinfection, and these rates were not affected by the addition of vitamin D3. These findings indicate that apoptosis is unlikely to be involved in the induction of antimycobacterial activity by D3.
      The second principal conclusion drawn from this study is that PI 3-K regulates the antimycobacterial action of D3. Several lines of evidence support this argument including the following: (i) PI 3-K is activated by D3 in THP-1 cells (Fig. 1 B; see Ref.
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ); (ii) D3-induced antimycobacterial activity is abrogated by PI 3-K inhibitors LY and Wm in both THP-1 cells and MDM independent of any cytotoxic effect on the cells (Fig. 2, Band C); (iii) D3-induced antimycobacterial activity is abrogated by antisense mRNA to class I PI 3-K (Fig.3 B); (iv) O⨪2 production by M. tuberculosis-infected THP-1 cells and MDM in response to D3 is inhibited by PI 3-K inhibitors LY and Wm (Fig. 5,A and C); (v) D3-induced O⨪2production by M. tuberculosis-infected THP-1 cells is inhibited by antisense mRNA to class I PI 3-K (Fig.5 B); and (vi) pretreatment of THP-1 cells with the PI 3-K inhibitors LY or Wm prevents p47phox and p67phoxtranslocation to the membrane fraction upon D3 treatment (Fig. 8, C and D).
      The finding that the antimycobacterial action of D3 is regulated by PI 3-K represents another important example of nongenomic signaling by this and other steroid hormones. The long standing paradigm for D3 action has been recognized to involve genomic signaling in which hormone binds to the VDR. The D3·VDR complex then translocates to the nucleus, where the ligand-activated transcription factor VDR directly regulates transcription by binding to vitamin D response elements in D3-activated genes (
      • Malloy P.J.
      • Feldman D.
      ). However, numerous reports have shown that D3 also acts via nongenomic signaling, where cellular responses are brought about independent of de novotranscription from a classical vitamin D response element (
      • Bhatia M.
      • Kirkland J.B.
      • Meckling-Gill K.A.
      ,
      • Berry D.M.
      • Antochi R.
      • Bhatia M.
      • Meckling-Gill K.A.
      ,
      • Darwish H.M.
      • DeLuca H.F.
      ). Moreover, it was recently shown that D3 treatment of human monocytes results in the rapid activation of PI 3-K leading to monocyte differentiation (
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ). In the latter report, a novel nongenomic mechanism of action of D3 was shown to be regulated by a VDR·PI 3-K signaling complex. Similar findings were also recently reported for signaling by estrogen, where hormone treatment induced the formation of a signaling complex including the estrogen receptor-α and the p85α regulatory subunit of PI 3-K (
      • Simoncini T.
      • Hafezi-Moghadam A.
      • Brazil D.P.
      • Ley K.
      • Chin W.W.
      • Liao J.K.
      ).
      PI 3-K has been shown to regulate a wide range of physiological processes including movement of organelle membranes, cytoskeletal rearrangement, cell proliferation, apoptosis, etc. (
      • Toker A.
      • Cantley L.G.
      ,
      • Franke T.F.
      • Kaplan D.R.
      • Cantley L.C.
      ). Importantly, PI 3-K has also been shown to regulate the oxidative burst. For example, PI 3-K has been shown to be required for triggering the NADPH oxidase in response to particulate stimuli through activating Fcα and Fcγ receptors in neutrophils (
      • Lang M.L.
      • Kerr M.A.
      ). Similarly, based upon effects of the PI 3-K inhibitor wortmannin, a role for this enzyme in regulating the production of superoxide by human neutrophils in response to soluble stimuli such as fMLP (
      • Santoro P.
      • Cacciapuoti C.
      • Palumbo A.
      • Graziano D.
      • Annunziata S.
      • Capasso L.
      • Formisano S.
      • Ciccimarra F.
      ) and PMA (
      • Yang M.
      • Wu W.
      • Mirocha C.J.
      ) has also been suggested. In the present study, PI 3-K was found to be required to trigger the oxidative burst in response to D3 in M. tuberculosis-infected cells. However, it appears that activation of PI 3-kinase alone is insufficient to bring about full oxidase activation, since treatment of THP-1 cells with D3 alone activates PI 3-K (Fig.1 B, lanes 6, 8, and10 versus lanes 5,7, and 9; see Ref.
      • Hmama Z.
      • Nandan D.
      • Sly L.
      • Knutson K.L.
      • Herrera-Velit P.
      • Reiner N.E.
      ), but at best this elicits only modest superoxide production (Fig. 5, AC). In contrast, prior infection of cells with live M. tuberculosis primed cells for an enhanced D3-induced oxidative burst, and this was not recapitulated by either latex beads or killed M. tuberculosis(Fig. 6). Consistent with these findings, only in cells infected with live bacilli did D3 induce translocation of the oxidase components p47phox and p67phox to the membrane fraction for oxidase assembly (Fig. 8). These findings suggest that liveM. tuberculosis or some factor unique to the phagosome containing viable M. tuberculosispotentiates phagocytes to undergo a vigorous oxidative burst in response to D3.
      In summary, this report identifies a key mechanism of D3-induced host defense against tuberculosis and a key regulatory pathway required to bring about this effector mechanism. The findings demonstrate that the antimycobacterial activity of D3 in human macrophages is due to activation of the NADPH oxidase and the production of reactive oxygen intermediates. Furthermore, they show that PI 3-K is necessary but not sufficient for assembly of a functional oxidase in response to D3 inM. tuberculosis-infected cells. This represents another novel example of nongenomic signaling by vitamin D3. Ongoing studies concerned with the basis forM. tuberculosis priming and the additional regulatory pathways involved should provide additional insight into host resistance to mycobacterial disease and the associated inflammatory consequences.

      REFERENCES

        • Kaufmann S.H.
        • van Embden J.D.
        Trends Microbiol. 1993; 1: 2-5
        • Day M.
        New Sci. 1998; 2127: 21
        • Armstrong J.A.
        • Hart P.D.
        J. Exp. Med. 1975; 142: 1-16
        • Hingley-Wilson S.M.
        • Sly L.M.
        • Reiner N.E.
        • McMaster W.R.
        Mod. Aspects Immunobiol. 2000; 1: 96-101
        • Malloy P.J.
        • Feldman D.
        Am. J. Med. 1999; 106: 355-370
        • Hmama Z.
        • Nandan D.
        • Sly L.
        • Knutson K.L.
        • Herrera-Velit P.
        • Reiner N.E.
        J. Exp. Med. 1999; 190: 1583-1594
        • Gniadecki R.
        Biochem. Pharmacol. 1998; 56: 1273-1277
        • Janis A.E.
        • Kaufmann S.H.E.
        • Schwartz R.H.
        • Pardoll D.M.
        Science. 1989; 244: 713-716
        • Phillips W.
        • Hamilton J.A.
        J. Immunol. 1989; 142: 2445-2449
        • Duncan R.
        • McConkey E.H.
        Eur. J. Biochem. 1982; 123: 535-538
        • Manolagas S.C.
        • Hustmyer F.G.
        • Yu X.P.
        Kidney Int. Suppl. 1990; 29: 9-16
        • Abe E.
        • Miyaura C.
        • Sakagami H.
        • Takeda M.
        • Konno K.
        • Yamazaki T.
        • Yoshiki S.
        • Suda T.
        Proc. Natl. Acad. Sci. U. S. A. 1981; 78: 4990-4994
        • Tanaka H.
        • Abe E.
        • Miyaura C.
        • Kuribayashi T.
        • Konno K.
        • Nishii Y.
        • Suda T.
        Biochem. J. 1982; 204: 713-719
        • Kreutz M.
        • Andreesen R.
        Blood. 1990; 76: 2457-2461
        • Schwende H.
        • Fitzke E.
        • Ambs P.
        • Dieter P.
        J. Leukocyte Biol. 1996; 59: 555-561
        • Zhang D.-E.
        • Hetherington C.J.
        • Gonzalez D.A.
        • Chen H.-M.
        • Tenen D.G.
        J. Immunol. 1994; 153: 3276-3285
        • Davies P.D.
        Tubercle. 1985; 66: 301-306
        • Roy S.
        • Frodsham A.
        • Saha B.
        • Hazra S.K.
        • Mascie-Taylor C.G.
        • Hill A.V.
        J. Infect. Dis. 1999; 179: 187-191
        • Wilkinson R.J.
        • Llewelyn M.
        • Toossi Z.
        • Patel P.
        • Pasvol G.
        • Lalvani A.
        • Wright D.
        • Latif M.
        • Davidson R.N.
        Lancet. 2000; 355: 618-621
        • Crowle A.J.
        • Ross E.J.
        • May M.H.
        Infect. Immun. 1987; 55: 2945-2950
        • Rook G.A.
        • Steele J.
        • Fraher L.
        • Barker S.
        • Karmali R.
        • O'Riordan J.
        • Stanford J.
        Immunology. 1986; 57: 159-163
        • Fry M.J.
        Biochim. Biophys. Acta Mol. Basis Dis. 1994; 1226: 237-268
        • Toker A.
        • Cantley L.G.
        Nature. 1997; 387: 673-676
        • Fukui Y.
        • Ihara S.
        • Nagata S.
        J Biochem. (Tokyo). 1998; 124: 1-7
        • Allen L.A.
        • DeLeo F.R.
        • Gallois A.
        • Toyoshima S.
        • Suzuki K.
        • Nauseef W.M.
        Blood. 1999; 93: 3521-3530
        • Stokes R.W.
        • Doxsee D.
        Cell. Immunol. 1999; 197: 1-9
        • Liu M.K.
        • Herrera-Velit P.
        • Brownsey R.W.
        • Reiner N.E.
        J. Immunol. 1994; 153: 2642-2652
        • Hmama Z.
        • Gabathuler R.
        • Jefferies W.A.
        • de Jong G.
        • Reiner N.E.
        J. Immunol. 1998; 161: 4882-4893
        • Volinia S.
        • Hiles I.
        • Ormondroyd E.
        • Nizetic D.
        • Antonacci R.
        • Rocchi M.
        • Waterfield M.D.
        Genomics. 1994; 24: 472-477
        • DeLeo F.R.
        • Jutila M.A.
        • Quinn M.T.
        J. Immunol. Methods. 1996; 198: 35-49
        • Laemmli U.K.
        FEMS Microbiol. Rev. 1970; 227: 680-685
        • Nandan D.
        • Reiner N.E.
        Infect. Immun. 1995; 63: 4495-4500
        • Clark R.A.
        • Volpp B.D.
        • Leidal K.G.
        • Nauseef W.M.
        J. Clin. Invest. 1990; 85: 714-721
        • DeLeo F.R.
        • Allen L.A.
        • Apicella M.
        • Nauseef W.M.
        J. Immunol. 1999; 163: 6732-6740
        • Bellamy R.
        • Ruwende C.
        • Corrah T.
        • McAdam K.P.
        • Thursz M.
        • Whittle H.C.
        • Hill A.V.
        J. Infect. Dis. 1999; 179: 721-724
        • Vazquez-Torres A.
        • Jones-Carson J.
        • Mastroeni P.
        • Ischiropolous H.
        • Fang F.C.
        J. Exp. Med. 2000; 192: 227-236
        • Babior B.M.
        Blood. 1999; 93: 1464-1476
        • Ohga S.
        • Ikeuchi K.
        • Kadoya R.
        • Okada K.
        • Miyazaki C.
        • Suita S.
        • Ueda K.
        J. Infect. 1997; 34: 147-150
        • Allen D.M.
        • Chng H.H.
        J. Infect. 1993; 26: 83-86
        • Chusid M.J.
        • Parrillo J.E.
        • Fauci A.S.
        JAMA. 1975; 233: 1295-1296
        • Gonzalez B.
        • Moreno S.
        • Burdach R.
        • Valenzuela M.T.
        • Henriquez A.
        • Ramos M.I.
        • Sorensen R.U.
        Pediatr. Infect. Dis. J. 1989; 8: 201-206
        • Cohen M.S.
        • Mesler D.E.
        • Snipes R.G.
        • Gray T.K.
        J. Immunol. 1986; 136: 1049-1053
        • Brett S.J.
        • Butler R.
        Clin. Exp. Immunol. 1988; 71: 32-38
        • Gordon A.H.
        • Hart P.D.
        Infect. Immun. 1994; 62: 4650-4651
        • Reiner N.E.
        Immunol. Today. 1994; 15: 374-381
        • Knutson K.L.
        • Hmama Z.
        • Herrera-Velit P.
        • Rochford R.
        • Reiner N.E.
        J. Biol. Chem. 1998; 273: 645-652
        • Maiti D.
        • Bhattacharyya A.
        • Basu J.
        J. Biol. Chem. 2000; 276: 329-333
        • Chan J.
        • Xing Y.
        • Magliozzo R.S.
        • Bloom B.R.
        J. Exp. Med. 1992; 175: 1111-1122
        • MacMicking J.D.
        • North R.J.
        • LaCourse R.
        • Mudgett J.S.
        • Shah S.K.
        • Nathan C.F.
        Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 5243-5248
        • Thoma-Uszynski S.
        • Stenger S.
        • Takeuchi O.
        • Ochoa M.T.
        • Engele M.
        • Sieling P.A.
        • Barnes P.F.
        • Rollinghoff M.
        • Bolcskei P.L.
        • Wagner M.
        • Akira S.
        • Norgard M.V.
        • Belisle J.T.
        • Godowski P.J.
        • Bloom B.R.
        • Modlin R.L.
        Science. 2001; 291: 1544-1547
        • Arbibe L.
        • Mira J.-P.
        • Teusch N.
        • Kline L.
        • Mausumee G.
        • Mackman N.
        • Godowski P.J.
        • Ulevitch R.J.
        • Knaus U.G.
        Nat. Immunol. 2001; 1: 533-540
        • Santucci M.B.
        • Amicosante M.
        • Cicconi R.
        • Montesano C.
        • Casarini M.
        • Giosuè S.
        • Bisetti A.
        • Colizzi V.
        • Fraziano M.
        J. Infect. Dis. 2000; 181: 1506-1509
        • Kremer L.
        • Estaquier J.
        • Brandt E.
        • Ameisen J.C.
        • Locht C.
        Eur. J. Immunol. 1997; 27: 2450-2456
        • Placido R.
        • Mancino G.
        • Amendola A.
        • Mariani F.
        • Vendetti S.
        • Piacentini M.
        • Sanduzzi A.
        • Bocchino M.L.
        • Zembala M.
        • Colizzi V.
        J. Pathol. 1997; 181: 31-38
        • Durrbaum-Landmann I.
        • Gercken J.
        • Flad H.D.
        • Ernst M.
        Infect. Immun. 1996; 64: 5384-5389
        • Keane J.
        • Remold H.G.
        • Kornfeld H.
        J. Immunol. 2000; 164: 2016-2020
        • Klingler K.
        • Tchou-Wong K.M.
        • Brandli O.
        • Aston C.
        • Kim R.
        • Chi C.
        • Rom W.N.
        Infect. Immun. 1997; 65: 5272-5278
        • Bhatia M.
        • Kirkland J.B.
        • Meckling-Gill K.A.
        J. Biol. Chem. 1995; 270: 15962-15965
        • Berry D.M.
        • Antochi R.
        • Bhatia M.
        • Meckling-Gill K.A.
        J. Biol. Chem. 1996; 271: 16090-16096
        • Darwish H.M.
        • DeLuca H.F.
        Prog. Nucleic Acid Res. Mol. Biol. 1996; 53: 321-344
        • Simoncini T.
        • Hafezi-Moghadam A.
        • Brazil D.P.
        • Ley K.
        • Chin W.W.
        • Liao J.K.
        Nature. 2000; 407: 538-541
        • Yang M.
        • Wu W.
        • Mirocha C.J.
        Immunopharmacol. Immunotoxicol. 1996; 18: 597-608
        • Franke T.F.
        • Kaplan D.R.
        • Cantley L.C.
        Cell. 1997; 88: 435-437
        • Lang M.L.
        • Kerr M.A.
        Biochem. Soc. Trans. 1997; 25: S603
        • Santoro P.
        • Cacciapuoti C.
        • Palumbo A.
        • Graziano D.
        • Annunziata S.
        • Capasso L.
        • Formisano S.
        • Ciccimarra F.
        Ital. J. Biochem. (Engl. Ed.). 1998; 47: 13-18