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Basic Science Aspects

Surfactant Proteins-A and -D Attenuate LPS-Induced Apoptosis in Primary Intestinal Epithelial Cells (IECs)

Zhang, Linlin∗,†; Meng, Qinghe; Yepuri, Natesh; Wang, Guirong; Xi, Xiuming; Cooney, Robert N.

Author Information
doi: 10.1097/SHK.0000000000000919

Abstract

INTRODUCTION

The surfactant proteins A (SP-A) and D (SP-D) are members of the C-type lectin (collectin) family which are especially important in pulmonary immunity and host defense (1, 2). SP-A and SP-D are hydrophilic proteins composed of different subunits including: the N-terminal domain, a collagenous-like triple helix and neck region, and the C-terminal carbohydrate recognition domain (1). SP-A and SP-D bind to monosaccharides with high, but slightly different affinities and interact with bacterial cell surfaces to act as opsonins. They also demonstrate direct bactericidal activity by altering cell membrane permeability (3). SP-A and SP-D are also involved in clearing foreign debris and apoptotic cells in many tissues and can regulate inflammatory responses through interactions with host pattern recognition receptors including: toll-like receptor 4 (TLR4), MD2, CD14, CR3 (CD11), and others (4). The expressions of SP-A and SP-D are not only found in lung, but also they are detected in extrapulmonary mucosal tissues such as stomach, intestine, and kidney (5).

Recent studies from our laboratory provide evidence SP-A and SP-D ameliorate lung injury in experimental pneumonia, but also appear to protect both gut and kidney from sepsis-induced organ injury (6, 7). These findings prompted us to further investigate the roles of endogenous and exogenous SP-A and SP-D in directly regulating intestinal injury and apoptosis. The current study focuses on the gut because of its central role in the progression of sepsis, systemic inflammation, and multiple organ dysfunction syndrome (MODS). Furthermore, many aspects of gut function are altered during sepsis including: gut integrity and barrier function, inflammatory cytokine production, and apoptosis of intestinal epithelial cells (8). Additionally, interventions that reduce intestinal epithelial apoptosis during sepsis are associated with improved survival (9).

The process of apoptotic cell death is characterized by preservation of membrane integrity, cell and organelle shrinkage, a unique pattern of DNA breakdown, the lack of associated inflammation, and immunosuppression. Lipopolysaccharide (LPS), the major component of the cell membrane from gram-negative bacteria, has been shown to trigger apoptosis in intestinal epithelia by binding to TRL4, activating mitogen-activated protein kinase (MAPK) and cleaved caspase-3 which activates pro-apoptotic pathways resulting in DNA degradation and cell death (10). The phosphorylation of Src homology region 2 domain-containing phosphatase-1 (SHP-1) is associated with the activation of MAPK (11). Apoptosis may be regulated by the activation of the MAPK signaling pathway or by the Bcl-2 family of intracellular proteins which inhibit apoptosis and/or the pro-apoptotic protein BAX (12). The ratio of BAX/Bcl-2 is commonly used to assess the “pro-apoptotic” intracellular environment. In contrast, SP-A and SP-D have been shown to act outside the cell to enhance the uptake of apoptotic cells by alveolar macrophages in vitro, to regulate host immune defenses and meditate inflammatory responses (13). However, the precise roles of SP-A and SP-D in directly regulating intestinal apoptosis and the P38 MAPK pathway are not well understood.

The current study examines the roles of SP-A, SP-D, and the P38 MAPK signaling pathway in regulating LPS-induced intestinal epithelial injury and apoptosis. Our experimental model uses primary IEC cultures from SP-A/D KO and wild-type (WT) mice followed by LPS stimulation with or without exogenous SP-A or SP-D and examines the effects on apoptosis and its mediators. The p-P38 MAPK agonist U46619 and the p-P38 MAPK kinase inhibitor SB203580 are used to delineate the role of p-P38 MAPK signaling. Our results provide evidence that SP-A and SP-D act as protective roles in LPS-induced caspase-3 cleavage, the BAX/Bcl-2 ratio, and apoptosis in primary IECs. They also show LPS-induced activation of P38 MAPK is attenuated by pretreatment with SP-A or SP-D. Although the exact mechanisms by which SP-A and SP-D protect IECs from apoptosis remain incompletely understood, our results provide evidence they attenuate the induction of LPS receptors TLR4 and CD14 and inhibit LPS-induced activation of the p-P38 MAPK pathway and its sequelae on the intracellular regulation of apoptosis.

METHODS AND MATERIALS

Animals

SP-A/D KO mice (C57BL/6 background) used in this study were gifted by Dr. Samuel Hawgood (University of California, San Francisco, Calif). SP-A/D KO mice were bred in the SUNY Upstate Medical University animal facility and genotypes were verified per animal use protocol, IACUC #270.

C57BL/6 WT mice were obtained from Charles River Laboratories (Wilmington, Mass) for a separate project by Dr Li-Ru Zhao and gut tissue was donated for this project according to SUNY Upstate Medical University animal use protocol, IACUC #338. All mice were housed in temperature-controlled rooms (22°C) on a 12 h light–dark schedule in our animal facility and allowed mouse chow and water ad libitum. All animal work was conducted at SUNY Upstate Medical University, approved by the institutional animal care committee and all experimental procedures on animals were carried out in accordance with the NIH and ARRIVE guidelines on the use of laboratory animals.

Primary IEC isolation and experiments

IECs were isolated using previously described methods (6, 14). Briefly, intestinal segments were harvested from euthanized mice, opened longitudinally, and washed in Ca2+ and Mg2+ free Hanks balanced salt solution (HBSS) (Gibco, Gaithersburg, Md) containing 2% glucose, 25 ng of amphotericin B per milliliter, 100 U of Penicillin-Streptomycin per milliliter (Gibco). Intestinal segments were minced into 1-mm fragments and incubated for 10 min at 37°C on a shaker platform in HBSS containing 60 U/mL of collagenase I (Sigma, St. Louis, Mo), 0.02 mg/mL of dispase I (Boehringer Mannheim, Indianapolis, Ind), 2% bovine serum albumin, and 0.2 mg/ml of soybean trypsin inhibitor (Sigma, St. Louis, Mo). Cells were centrifuged at 120 × g for 3 min in DMEM plus 2% sorbitol. Cell pellets containing intact crypts and sheets of epithelia were cultured on Matrigel (Collaborative Biomedical Products, Bedford, Mass) coated plates in defined media as previously described (6, 14). Cells were cultured in 5% CO2 at 37°C with periodic supplementation of medium. IEC cell growth and morphology were assessed by phase microscopy and Ber-EP4 (epithelial antigen) expression (> 80% cells stain positive for Ber-EP4, data not shown).

Pilot experiments were performed to determine the LPS dose used in this study by incubating 0, 10, 50, 100, 250, and 500 ug/mL of LPS (filtered through a 0.2 μ filter) for 24 h with our primary IEC cultures. Cell viability, cleaved caspase 3, BAX, and secreted IL-1β (Supplemental Data 1, http://links.lww.com/SHK/A597) in the media were measured. Cell viability (by MTT assay) was 80% or higher with 50 to 100 μg of LPS and decreased dramatically when LPS was increased to the 250 to 500 μg/mL dose range. LPS doses of 50 to 100 μg/mL were associated with increased levels of cleaved caspase 3 and BAX proteins as well as increased levels of IL-1 in the media. Based on these data, the dose of 100 μg/mL of LPS was chosen for subsequent studies. All IECs were inspected under the microscope before use in experiments to verify they appeared histologically “healthy.” Prior to treatment, IECs monolayers were washed with PBS three times. IECs from SP-A/D KO and WT mice were stimulated with LPS 100 μg/mL in serum-free media and harvested at 24 h. In experiments examining the role of exogenous surfactant, IECs from WT mice were pretreated with SP-A (10 μg/mL), SP-D (10 μg/mL), or vehicle for 20 h, then 100 μg/mL of LPS for 24 h. U46619, a P38 MAPK activator, was used in experiments to examine the inhibitory effect of exogenous surfactant on p-P38 MAPK activation. IECs from WT mice were pretreated with SP-A (10 μg/mL), SP-D (10 μg/mL) or vehicle for 20 h, then 1 μM of U46619 for 24 h. SB203580 (SB), a P38 MAPK inhibitor, was used to block P38 MAPK activation. IECs from WT mice were pretreated with SB203580 (40 μM) or vehicle for 20 h, then 100 μg/mL of LPS for 24 h.

Reagents

LPS was purchased from Sigma (cat. no. l3880, Louis, Mo). Native human SP-A was purified from bronchoalveolar lavage fluid (BALF) as previously described and purity of the SP-A preparation was verified by SDS-PAGE followed by silver staining. Recombinant human SP-D was purchased from Creative BioMart (can. no. SFTPD-3868H, Shirley, NY). The P38 MAPK agonist (U46619) was obtained from Santa Cruz Biotechnology (Dallas, Tex) and the P38 MAPK inhibitor (SB203580) was obtained from Sigma (St. Louis, Mo). Our sources for antibodies are as follows: Cleaved caspase-3 (1:1,000; cat. no. AB49822, Abcam, Cambridge, Mass), P38 MAPK (1:1,000; cat. no. #9212; Cell Signaling Technology, Danvers, Mass), p-P38 MAPK (1:1,000; cat. no. #9211; Cell Signaling Technology, Danvers, Mass), TLR4 (1:200; cat. no. SC-10741; Santa Cruz Biotechnology, Dallas, Tex), CD14 (1:200; cat. no. SC-9150; Santa Cruz Biotechnology, Dallas, Tex), SHP-1 (1:2,000; cat. no. ab131537, Abcam, Cambridge, Mass), and p-SHP-1 (1:2,000; cat. no. ab131500, Abcam, Cambridge, Mass).

Western blot analyses

IECs were homogenized and lysed on ice using protein lysis buffer including a cocktail of protease inhibitors (Roche, Indianapolis, Ind) and cell lysates were centrifuged at 12,000 rpm at 4°C for 5 min. The supernatant was collected for the Western blot to detect protein expression. Protein concentrations were assayed using BCA protein kit (Thermo Scientific, Rockford, Ill). Protein samples (20 μg) were denatured with sample loading buffer at 95° for 5 min, subjected to 12% SDS-PAGE gel for 2 h at 100 V then transferred to a PVDF membrane for 2 h at 60 V. Membranes were blocked in 5% nonfat dry milk in Tris-buffered saline (TBS) with 0.1% Tween-20 at room temperature for 1 h. Membranes were incubated at 4°C overnight with primary antibodies (cleaved caspase-3, BAX, Bcl-2, TLR4, t-P38, p-P38, SHP-1, p-SHP-1). The following day, membranes were rinsed and then incubated for 1 h at room temperature with secondary Horseradish Peroxidase (HRP)-conjugated antibody (Bio-Rad, Hercules, Calif). Protein bands were visualized using Pierce ECL Western Blotting detection solution (Thermo Scientific, Rockford, Ill) according to the manufacturer's instructions and recorded by exposure of the membrane to x-ray film. The membranes were stripped and probed with GAPDH antibody (Santa Cruz Biotechnology, Santa Cruz, Calif) to verify equal protein loading. Band intensity was quantified by the use of Quantity One software (Bio-Rad, Hercules, Calif). Western blot results were reported as relative densitometry units (RDU) normalized to GAPDH or β-actin.

TUNEL assay

A commercially available TUNEL assay (Life Technologies Co, Ore) was used to detect apoptotic IECs in primary cell culture according to the manufacturer's instructions. Briefly, media was removed from IECs seeded in 96-well plates, rinsed with PBS, then fixed with 4% paraformaldehyde for 15 min. 100 μL of 0.25% Triton X-100 was added for permeabilization and incubated for 20 min at room temperature, after removing the fixative solution. Then 100 μL of TdT reaction buffer was supplied and incubated for 10 min at 37°C. 50 μL of TdT reaction mixture was then added and incubated for 60 min at 37°C after removing TdT reaction buffer. Immediately 50 μL of the Click-iT Plus TUNEL reaction cocktail was added after washing and incubated for 30 min at 37°C, protected from light. In the end, the nuclei labeling with DAPI was followed. Apoptotic cells were revealed by Nikon Eclipse TE 2000-Umicroscope (Nikon Instruments Inc, Melville, NY). Apoptotic cells were quantified by counting TUNEL-positive cells from five randomly selected consecutive fields at ×400 magnification by two experienced investigators in a blinded manner. Apoptotic index was calculated as the number of TUNEL-positive cells expressed as percentage of total cells.

Supplemental immunocytochemistry (ICC) and immunofluorescence (IF) data

ICC was also used to measure cleaved caspase 3. The IECs were seeded in a 96-well plate. First, the medium was removed gently, then 100 μL of 4% paraformaldehyde solution (USB Co, Cleveland, Ohio) was added to attach cultured cells, incubated for 10 min at room temperature. Next, 4% PFA was removed and 100 μL per well of 0.5% Triton-100 (Sigma, St. Louis, Mo) in PBS was added for permeabilization, incubated for 5 min at room temperature. Cells are gently rinsed three times with PBS after removing permeabilization solution and then 100 μL of blocking buffer was added and incubated for 1 h at room temperature. Primary antibody solution, cleaved caspase-3 (1:400; cat. no. #9664; Cell Signaling Technology, Danvers, Mass) was added and incubated overnight at 4°C after removing blocking buffer. The next day, primary antibody solution was removed and gently rinsed three times with PBS. Secondary antibody solution donkey anti-rabbit IgG (1: 200; cat no. ab150073; Abcam, Cambridge, Mass) was added and incubated for 1 h in a dark environment. DAB was supplied after the secondary antibody solution was removed. Cells were imaged with Nikon Eclipse TE 2000-Umicroscope (Nikon Instruments Inc, Melville, NY).

IF was also used to measure Bcl-2 and BAX. Cellular localizations were carried out in 96-well plates seeded with IECs. After fixation with paraformaldehyde solution, 100 μL per well of 0.5% Triton-100 (Sigma, St. Louis, Mo) in PBS was added for permeabilization, incubated for 5 min at room temperature, and then blocked with normal donkey serum at room temperature for 1 h. IECs were incubated with primary antibody overnight at 4°C, and then with second primary antibodies for 1 h after blocking. Primary antibodies used were BAX (1:50; cat. no. SC-526; Santa Cruz Biotechnology, Dallas, Tex), Bcl-2 (1:50; cat. no.SC-526; Santa Cruz Biotechnology, Dallas, Tex). Secondary antibodies were donkey anti-rabbit IgG (1: 200; cat no. ab150073; Abcam, Cambridge, Mass), and donkey anti-goat IgG (1:100; Cat. no. A-11058; Thermo Fisher, Rockford, Ill). Cells were gently rinsed three times with PBS after removing secondary antibody solution and. In the end, Fluoroshield mounting medium with DAPI (Cat. no. 104139; Abcam) was supplied and cells were imaged with Nikon Eclipse TE 2000-Umicroscope (Nikon Instruments Inc, Melville, NY).

STATISTICAL METHODS

Data are presented as means ± SEM. Statistical analysis of the data was performed using Student t test or ANOVA followed by Student–Newman–Keuls post testing using Prism 5.0 (GraphPad Software, San Diego, Calif). Differences among groups (n = 5–8/group) were considered significant at P ≤ 0.05.

RESULTS

SP-A and SP-D regulate LPS-induced apoptosis

IECs from SP-A/D KO and WT mice were treated with LPS. Apoptotic index, cleaved caspase-3 levels, and the BAX/Bcl-2 ratio were measured (Fig. 1). Incubation with LPS significantly increased the apoptotic index, cleaved caspase-3 levels, the relative abundance of BAX and BAX/Bcl2 ratio in WT IECs. LPS-treated IECs from SP-A/D KO mice demonstrate significantly more apoptosis and increased expression of cleaved caspase-3, BAX and the BAX/Bcl-2 ratio compared to unstimulated controls and LPS-stimulated WT IECs. To further characterize the regulation of apoptosis by SP-A and -D, IECs from WT mice were pretreated with vehicle, SP-A, or SP-D and then stimulated with LPS. As noted above, LPS stimulation significantly increased apoptotic index, cleaved caspase-3 and BAX, as well as the BAX/Bcl2 ratio in WT IECs (Fig. 2, P < 0.05 vs. Control). Pretreatment with SP-A or SP-D significantly attenuated LPS-induced increases in apoptosis and the associated changes in cleaved caspase-3, BAX, and the BAX/Bcl-2 ratio. Cleaved caspase 3 was also measured by ICC (positive staining: brown color), BAX and Bcl-2 were also measured by IF assay (positive staining: green; merged staining: white). Western blot, ICC, and IF techniques provided similar results (Supplemental Data 2, http://links.lww.com/SHK/A598, & 3, http://links.lww.com/SHK/A599).

Fig. 1
Fig. 1:
Endogenous SP-A and SP-D regulate LPS-induced apoptosis.
Fig. 2
Fig. 2:
Exogenous SP-A and SP-D regulate LPS-induced apoptosis.

SP-A and SP-D regulate LPS-induced P38 MAPK activation

Activation of the P38 MAPK signaling pathway by LPS has been implicated in the pathogenesis of apoptosis in IECs by several studies. P38 MAPK is activated as a result of tyrosine phosphorylation by MKK3/6 and inactivated by MAPK phosphatase-1 (MKP-1)-mediated dephosphorylation. The ratio of phosphorylated to total P38 MAPK (p-P38 MAPK/t-P38 MAPK) is generally viewed as an indicator of P38 MAPK activation. As shown in Figure 3A, LPS stimulation increased the ratio of p-P38 MAPK/t-P38 MAPK in IECs from both WT and SP-A/D KO mice (P < 0.05 vs. Controls). However, LPS-induced activation of P38 MAPK was significantly increased in IECs from SP A/D KO mice compared with the WT LPS group. The effects of SP-A and SP-D on LPS-induced activation of P38 MAPK are shown in Figure 3B. Pretreatment of WT IECs with either SP-A or SP-D significantly attenuated the LPS-induced activation of P38 MAPK (P < 0.05 vs. LPS alone).

Fig. 3
Fig. 3:
Regulation of P38 MAPK signaling by LPS, SP-A, and SP-D.

Regulation of P38 MAPK signaling by LPS, SP-A, and SP-D

To further characterize the effects of SP-A and SP-D on LPS-induced P38 MAPK signaling and apoptosis, we examined their effects when used in conjunction with the MAPK agonist (U46619, a thromboxane A2 mimetic) and the P38 MAPK inhibitor (SB203580, which reduces epirubicin-induced cell injury and caspase-3/7 activity). As shown in Figure 4, treatment with U46619 significantly increased the apoptotic index, cleaved caspase-3 levels, and relative abundance of BAX and BAX/Bcl2 ratio in WT IECs. Pretreatment with either SP-A or SP-D significantly reduced the induction of cleaved caspase-3, BAX and the BAX/Bcl-2 ration as well as apoptosis in U46619-treated cells. Next, we examined the effects of SB203580, SP-A, and SP-D on LPS-induced apoptosis in WT IECs. The P38 MAPK inhibitor SB203580 attenuated LPS-induced apoptosis and apoptotic mediators (cleaved caspase-3, BAX, and BAX/Bcl-2 ratio), while the addition of either SP-A or SP-D to SB203580 restored apoptosis to control levels (Fig. 5). In U46619 and SB203580-treated cells, cleaved caspase 3 was also measured by ICC (positive staining: brown color), BAX and Bcl-2 were also measured by IF assay (positive staining: green; merged staining: white). Western blot, ICC and IF techniques provided similar (Supplemental Data 4, http://links.lww.com/SHK/A600, & 5, http://links.lww.com/SHK/A601).

Fig. 4
Fig. 4:
Regulation of IECs apoptosis by P38 MAPK agonist U46619, SP-A, and SP-D.
Fig. 5
Fig. 5:
Regulation of apoptosis by P38 MAPK inhibitor SB203580, SP-A, and SP-D.

SP-A and SP-D regulate LPS-induced expressions of TLR4 and CD14

The cellular effects of LPS are mediated by Toll-like receptor 4 (TLR4) protein and multiple associated proteins including: LPS binding protein (LBP), CD14, and MD2. MD2 and CD14 are small glycoproteins that can interact with both LPS and TLR4 to activate or inhibit the TLR4 signaling pathway. Furthermore, SP-A and SP-D have been shown to inhibit LPS-mediated TLR4 activation by multiple mechanisms including: interactions with LPS and binding to TLR4, CD14 and MD-2. To determine if the inhibitory effects of SP-A or SP-D were mediated by changes in the expression of the LPS receptor or associated proteins, we measured the levels of TLR4 and CD14 in IECs from WT and SP-A/D KO mice. As shown in Figure 6A and B, LPS significantly increased the relative abundance of TLR4 and CD14 in IECs from both WT and SP-A/D KO mice. Nevertheless, the relative abundance of both TLR4 and CD14 was increased in LPS-treated IECs from SP-A/D KO mice. This finding was supported by the ability of pretreatment with either SP-A or SP-D to attenuate the LPS-induced increase in TLR4 and CD14 in WT IECs (Fig. 6, C and D).

Fig. 6
Fig. 6:
SP-A and SP-D regulate LPS-induced expressions of TLR4 and CD14.

SP-A and SP-D regulate LPS-induced expressions of Src SHP-1

Tyrosine phosphorylation and dephosphorylation of proteins is a key regulatory system of signal transduction which controls many aspects of cellular functions including cell growth, differentiation, and mitotic cycle. SHP-1, also called tyrosine-protein phosphatase non-receptor type 6, is a well-known signaling molecule that regulates a variety of cellular processes (15). In order to examine the effect of SP-A and SP-D on phosphorylation of SHP-1, total SHP-1 (t-SHP-1) and phosphorylated SHP-1 (p-SHP-1) were detected as shown in Figure 7. LPS significantly increased the relative abundance of p-SHP-1 but not t-SHP-1 in cells from WT mice. Pretreatment with SP-A and SP-D attenuates the LPS-induced increase in p-SHP-1/t SHP-1. These provide evidence for SP-A and SP-D in regulating the phosphorylation of SHP-1.

Fig. 7
Fig. 7:
Effects of SP-A and SP-D on LPS-induced SHP-1 phosphorylation.

DISCUSSION

Multiple organ dysfunction (MODS) and failure (MOF) represent the leading cause of death in non-coronary ICUs and the most significant “potentially preventable” cause of death in multiple trauma patients surviving the golden hour (16). Despite our improved understanding of “risk factors” and clinical interventions to reduce organ failure, patients with sepsis and shock continue to suffer major morbidity related to MODS. From a temporal perspective, lung injury is seen in > 90% of patients with MODS and frequently precedes the subsequent deterioration of function in other organs. Furthermore, the severity of lung injury correlates with both the number and severity of other organ systems involved in patients with MOF (17, 18).

Although lung is frequently the first organ to “fail,” several lines of evidence implicate the gut in the pathogenesis of sepsis and shock-related MOF, especially as it relates to acute lung injury. Systemic infection and shock-related gut ischemia/reperfusion result in intestinal inflammation, gut barrier dysfunction, release of toxic mediators in mesenteric lymph, and apoptosis of intestinal epithelial cells (19). While the relative importance of proteolytic injury to the gut mucosa, inflammatory mediators in mesenteric lymph and intestinal epithelial apoptosis remains controversial, the gut clearly seems to play a central role in the pathogenesis of MOF. In addition, preventing intestinal epithelial apoptosis improves survival in a murine model combining radiation injury with pneumonia (20). With this in mind, the current study examines the role of SP-A and SP-D in the pathogenesis of LPS-induced intestinal epithelial cell apoptosis. Previous work from our laboratory in SP-A/D KO mice provides evidence SP-A and SP-D regulate sepsis-induced injury to lung, kidney, and intestine (6, 7). While the lung is viewed as the predominant source of circulating surfactant, both kidney and gut express SP-A and SP-D and the relative importance of local surfactant production in preventing organ injury in vivo is poorly understood. Our study provides the first direct evidence SP-A and SP-D can protect primary IECs from LPS-induced and P38 MAPK-induced apoptosis.

Homeostasis of the gut epithelium is maintained by an appropriate balance between cellular proliferation and death. Apoptotic cell death of IECs may be triggered by a variety of stressors including oxygen-free radicals, inflammatory cytokines, and/or LPS and appears to involve the activation of the TLR4 and P38 MAPK signaling pathways (21). While in vivo studies implicate SP-A and SP-D in the removal of apoptotic cells by alveolar macrophages (22), our study design (primary IECs cultures) and the reductions in cleaved caspase-3 levels and the BAX/Bcl-2 ratio suggest SP-A and SP-D act by attenuating or blocking the LPS-induced apoptotic stimuli in primary IECs. Our findings are consistent with other studies showing exogenous surfactant administration decreases lung injury and apoptosis by reducing cleaved caspase-3 (23).

The induction of apoptotic cell death by LPS involves signaling by both the TLR4 and P38 MAPK pathways (12, 24). As collectins, SP-A and SP-D can directly interact with TLR4 and CD14 through their carbohydrate recognition domains and interfere with the interactions between LPS, CD14, and the TLR4 receptor required for signaling, thus attenuating LPS-induced inflammation and apoptosis (4, 25–27). Although our experiments do not specifically test this mechanism, our results showing pretreatment of IECs with SP-A or SP-D attenuate apoptosis and its regulators (cleaved caspase-3, BAX, and BAX/Bcl-2 ratio) and reduce LPS-induced P38 MAPK activation could be explained in part by SP-A and SP-D binding to LPS, CD14, and TLR4. Studies showing the natural surfactant product Survanta is able to inhibit pro-inflammatory cytokine secretion and MAPK activation in alveolar macrophages are consistent with our findings. The ability of SP-D to attenuate LPS-induced inflammation in a human intestinal cell line (INT407) which overexpresses TLR4 is also consistent with this mechanism of action as well (28).

We believe we are the first to report that SP-A or SP-D can attenuate the induction of apoptosis and its mediators (cleaved caspase-3, BAX, and BAX/Bcl-2 ratio) by U46619, a P38 MAPK activator in WT IECs. These results suggest the presence of other mechanisms by which SP-A and SP-D regulate apoptotic cell death. Several potential cellular receptors for SP-A and SP-D have been identified including C1q or calreticulin that interacts with CD91 and more recently signal inhibitory regulatory protein α (SIRPα) (29–31). qRT-PCR data show C1q, calreticulin, and SIRPα are all expressed (data not shown) in our primary IEC cultures. Studies by Guo et al. provide evidence SP-A and/or SP-D can have pro- or anti-inflammatory effects depending on their nitrosylation status and which cellular receptor they bind. Thus, surfactant proteins -A and -D may also exert their protective effects on LPS-induced apoptosis of IECs through interactions with putative cell surface receptors like SIRPα. Consistent with these findings, SP-A or SP-D have inhibitory effects on LPS-induced apoptosis in WT IECs treated with SB203580 (a P38 MAPK inhibitor). Studies by Janssen et al. provide evidence SP-A and SP-D can attenuate P38 MAPK signaling in alveolar macrophages and activate the tyrosine phosphatase SHP-1 (30). Although SHP-1 does not appear to directly regulate P38 MAPK phosphorylation, it has been shown to indirectly regulate the MAPK pathway and LPS-induced IL-10 production in murine macrophages (11). Consistent with these studies our results demonstrate SP-A and -D can regulate LPS-induced phosphorylation of SHP-1. SP-A or SP-D mediated activation of mitogen-activated protein kinase phosphatase-1 (MAPK-1) which has been shown to prevent LPS-induced apoptosis in IECs represents another potential mechanism for our findings (32).

The increase in the relative abundance of TLR4 and CD14 proteins in LPS-treated IECs suggests their expression is regulated by the activation of TLR4 signaling (33). Consistent with this finding, the LPS-induced increase in CD14 and TLR4 was increased in IECs from SP-A/D KO mice and attenuated by pretreatment with either SP-A or SP-D. These data provide evidence endogenous and exogenous SP-A and SP-D can regulate the induction of CD14 and TLR4 by LPS in IECs.

Collectively, our findings provide novel evidence SP-A and SP-D can directly regulate LPS-mediated intestinal epithelial apoptosis. In summary SP-A and SP-D directly regulate LPS-induced apoptosis in IECs; SP-A and SP-D can attenuate apoptosis in IECs by LPS-induced and “direct” activation of the P38 MAPK signaling pathway; SP-A and SP-D can also attenuate the LPS-mediated increase in CD14 and TLR4 in IECs. The protective effects of SP-A and SP-D on LPS-induced may be due in part to their ability to directly bind with LPS, CD14, and TLR4. However, it is also possible the effects of SP-A/D are mediated by interactions with their cellular receptors calreticulin or SIRPα which regulate P38 MAPK activity and/or apoptosis. Our study has several limitations including: the activation status of p38 MAP kinase was only examined at one time point and as such is a “snapshot” of the activation/deactivation process which occurs over time; several in vivo studies suggest intestinal epithelial apoptosis is deleterious to the host and while our cell culture model is useful in elucidating the role of SP-A and SP-D in regulating this process they do not provide any information on whether this is beneficial or deleterious to the host; while we have identified several potential mechanisms for the effects of SP-A and SP-D on LPS-induced apoptosis, the relative importance of direct effects on LPS-binding proteins versus surfactant receptor-mediated events remains unclear and additional studies will be required to better understand the mechanisms by which SP-A and SP-D attenuate LPS-induced apoptosis in IECs. While SIRPα is expressed in primary IECs and LPS-induced phosphorylation of SHP-1 is attenuated by SP-A and SP-D, further studies are needed to delineate the specific roles of SIRPα and SHP-1 in regulating apoptosis in IECs.

Acknowledgment

The authors thank Dr S. Hawgood from the University of California, San Francisco, CA, for kindly providing SP-A/D KO mice.

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Keywords:

Apoptosis; intestinal epithelial cells (IECs); P38 mitogen-activated protein kinases (P38 MAPK); surfactant proteins-A and -D

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