Neutrophils decrease Ca2+-associated proteins and cellular proliferation in hepatocytes without cellular damage. To mimic the interaction between infiltrating neutrophils and hepatocytes, liver-derived HepG2 cells were cocultured with human neutrophils (Figure 1, A–E). Cxcl1 mRNA was elevated in the HepG2 cells, similar to the response of hepatocytes to infiltrating neutrophils in vivo (Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/JCI171691DS1) (20, 21). Although neutrophils can damage hepatocytes (3, 4), HepG2 cell viability was similar in cocultured cells and controls not exposed to neutrophils (Figure 1, A, B, and D). In contrast, neutrophil viability dropped progressively unless neutrophils were cocultured with HepG2 cells (Figure 1, B, C, and E). These results are consistent with recent reports showing that neutrophils in circulation have a short half-life, but infiltrating neutrophils change their phenotype to live from several days to 2 weeks (22–24). Neutrophils did not damage HepG2 cells by other measures as well, including albumin levels and mitochondrial membrane potential in the cells and alanine transaminase (ALT) levels in the culture medium (Supplemental Figure 1, B–E). Similar results were observed in primary mouse hepatocytes cocultured with mouse neutrophils (Supplemental Figure 1, F–K). Viability of mouse neutrophils also was significantly improved when cocultured with the hepatocytes (Supplemental Figure 1G).

Neutrophils decrease Ca2+-related proteins in hepatocytes without causing cell death. (A–C) Representative images of HepG2 cells and neutrophils after 20 hours of culture, using double staining (calcein-AM for alive [green], ethidium homodimer-1 for dead [red]). HepG2 cells only (A), coculture (B), neutrophils only (C). (D and E) Graphs comparing time series of cell viability with and without coculture. (D) Viability of HepG2 cells does not decrease over time regardless of coculture. (E) Viability of neutrophils decreases over time unless they are cocultured. Differences between alone and coculture are represented by daggers, and changes over time are indicated by asterisks; 5 fields per coverslips were measured. (F and G) Representative images double labeled with EdU (green) and Hoechst 33342 (blue) of HepG2 cells alone (F) and cocultured with neutrophils (G) after 12 hours. (H) HepG2 cell proliferation (EdU-positive cells) is decreased by coculture with neutrophils; 5–6 fields were measured per coverslip. (I) Ca2+ signals in HepG2 cells are altered by neutrophils. Representative tracings of Fluo-4 fluorescence intensity of HepG2 cells alone or cocultured with neutrophils for 20 hours. Cells were stimulated with 20 μM ATP. (J) Ca2+ signaling in HepG2 cells as measured by the AUC upon ATP stimulation is diminished by neutrophils. Nine coverslips of HepG2 cells (362 total cells) and 8 coverslips of HepG2 cells cocultured with neutrophils (279 total cells) were analyzed. (K) Representative immunoblots and (L) quantitation of the blots show that ITPR1, ITPR2, ITPR3, and SERCA2 are decreased in HepG2 cells, but calnexin and SEC61B are not, after coculture with neutrophils for 1 hour. ITPR1, ITPR2, and ITPR3 were blotted onto different membranes because of their close molecular weights, and their ratios to GAPDH were measured separately. (M) Representative immunoblots and (N) quantitation of the blots shows that ITPR2 and SERCA2 are decreased in primary human hepatocytes (HHC) that are cocultured for 1 hour. The graph represents neutrophils from 5 healthy volunteers. In A–N, all are from 3 independent experiments. Data are mean ± SD. NS, not significant. **P < 0.01; ****P < 0.0001; †††P < 0.001; ††††P < 0.0001 by unpaired, 2-tailed Student’s t test. Scale bars: 50 μm.

Since hepatocyte proliferation is reduced in AAH (25–27), the potential role of neutrophils was examined. The number of 5-ethynyl-2′-deoxyuridine–positive (EdU-positive) HepG2 cells was significantly reduced when cocultured with neutrophils (Figure 1, F–H). Hepatocyte proliferation is regulated by Ca2+ signaling (28, 29), and ATP-induced Ca2+ signals in HepG2 cells were found to be significantly reduced by coculture with neutrophils (Figure 1, I and J). Moreover, examining expression of major proteins involved in intracellular Ca2+ signaling revealed that all 3 ITPR isoforms (ITPR1, ITPR2, and ITPR3) and the sarcoplasmic reticulum/ER Ca2+-ATPase 2 (SERCA2) pump were dramatically reduced (Figure 1, K and L). Conversely, the ER-resident proteins SEC61b and calnexin were unchanged. ITPR2, the most abundant ITPR isoform in hepatocytes (17), also was reduced in primary human and mouse hepatocytes cocultured with neutrophils for 1 hour (Figure 1, M and N, and Supplemental Figure 1, H and I) and SERCA2 was reduced in human hepatocytes (Figure 1M). Together, these results show that neutrophils interact with hepatocytes in a nondestructive manner to impair expression of Ca2+-handling proteins and Ca2+ signaling.

Neutrophils decrease ITPR2 in HepG2 cells via a reversible nontranscriptional mechanism. To understand the mechanism of the neutrophil-induced decrease in Ca2+ signaling proteins in hepatocytes, time-course experiments were performed focusing on ITPR2, because it is also the principal Ca2+ channel protein responsible for hepatocyte proliferation (28, 30, 31). ITPR2 protein levels in HepG2 cells decreased as early as 1 hour after coculture and this decrease persisted at 4 hours and 20 hours of coculture (Figure 2, A and B). Ca2+ signaling in HepG2 cells was similarly decreased 1 hour after coculture with neutrophils (Supplemental Figure 2, A and B). To test whether this reduction in ITPR2 is transcriptionally regulated, ITPR2 gene expression was measured, but ITPR2 mRNA was increased rather than decreased after 1 hour of coculture (Figure 2, C and D). To determine whether this reduction is reversible, cocultured neutrophils were removed, and then HepG2 cells recovered by monoculture. Under these conditions, ITPR2 levels in HepG2 cells rapidly recovered, as did Ca2+ signals (Figure 2, E–G). These data suggest neutrophil-induced loss of ITPR2 in HepG2 cells occurs through protein degradation rather than mRNA transcription. Indeed, longer exposure of ITPR2 immunoblots of HepG2 cell lysates after neutrophil coculture showed a ladder pattern (Supplemental Figure 2C). Inhibitors of autophagy, proteasomes, and calpain were tested (32, 33), but none were able to prevent the decrease in ITPR2 (Figure 2, H and I). Inhibitors of trypsin and caspase 3 also failed to prevent ITPR2 reduction (Supplemental Figure 2, D and E) (34, 35). Collectively, these data suggest neutrophils reduce ITPR2 by a transient proteolytic mechanism.

The decrease in ITPR2 in HepG2 cells induced by neutrophils is rapid and reversible. (A) Representative immunoblots and (B) quantitation of ITPR2 levels in HepG2 cells cocultured with neutrophils show that loss of ITPR2 persists for up to 20 hours in the continued presence of neutrophils. (C and D) RT-qPCR shows that ITPR2 mRNA levels in HepG2 cells are increased after 1 hour and decreased after 20 hours of coculture with neutrophils. (E) Representative immunoblots and (F) quantitation show that ITPR2 levels begin to recover after neutrophils are removed. ITPR2 levels were measured in HepG2 cells cocultured with neutrophils for 20 hours, and then cells were washed to remove neutrophils and cultured for 2 more hours and collected (after neutrophil removal). Comparisons were relative to HepG2 cells cultured alone for 22 hours or cocultured with neutrophils. (G) Ca2+ signals in HepG2 cells progressively recover after neutrophils are removed. AUC of Fluo-4 fluorescence after stimulation with ATP (20 μM) was measured at 3 time points: after coculture with neutrophils for 1 hour (1 h, followed by complete removal of neutrophils for 1 hour [2 h] or 19 hours [20 h]). Data represent 5–7 coverslips each, with cell numbers n = 286, 208, 112. (H) Representative immunoblots and (I) quantitation shows that loss of ITPR2 in HepG2 cells is not blocked by treatment with MG132 (proteasome inhibitor, 50 μM), bafilomycin A1 (Baf, autophagy inhibitor, 50 nM), or Ac-Leu-Leu-Nle-aldehyde (ALLN, calpain inhibitor, 50 μM) for 1 hour, followed by coculture with neutrophils for 1 hour. Data are expressed as mean ± SD, n = 3–8. NS, not significant. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001 by unpaired, 2-tailed Student’s t test (B–D and F) or 1-way ANOVA with Tukey’s multiple-comparison test (G and I).

Direct contact with intact neutrophils is necessary for the decrease in ITPR2 in hepatocytes. Several factors secreted by neutrophils have been implicated in liver diseases (26, 36). However, ITPR2 was not decreased in HepG2 cells treated with neutrophil-conditioned medium (Figure 3, A and B) or in cells separated from neutrophils by a semipermeable membrane (Figure 3, C and D). These results suggest neutrophils require direct contact with HepG2 cells to reduce ITPR2. To evaluate this, glycosylphosphatidylinositol-anchored (GPI-anchored) plasma membrane proteins were stripped from neutrophils using phosphatidylinositol-specific phospholipase C (PI-PLC, 0.5 units/mL) to generate “naked” neutrophils (37). The effect of naked neutrophils on ITPR2 was significantly attenuated compared with untreated neutrophils (Figure 3, E and F). Bulk RNA-seq was used to compare global gene expression in HepG2 cells with and without neutrophil coculture, and Ingenuity Pathway Analysis (IPA) revealed that neutrophils altered signaling related to cell adhesion and diapedesis in HepG2 cells (Supplemental Figure 3A). Integrin isoforms α2 and αM (ITGA2 and ITGM) belong to the top 5 IPA signaling pathways of cocultured HepG2 cells and can act as binding partners for neutrophils on some epithelia (37–39). However, blocking antibodies directed against these integrins did not prevent reduction of ITPR2 in HepG2 cells cocultured with neutrophils (Figure 3, G and H). An additional mechanism by which neutrophils come into direct contact with their targets is neutrophil extracellular traps (NETs), which can occur in liver (40). Typical mediators of NETs include high mobility group box 1 protein (HMGB1), neutrophil elastase, and myeloperoxidase (MPO) (41). However, extracellular administration of HMGB1, neutrophil elastase, or MPO to HepG2 cells had no effect on ITPR2 (Figure 3, I and J). Coculture for 4 hours with neutrophils in which NETs were induced by phorbol 12-myristate 13-acetate (PMA, 200 nM) also reduced ITPR2 in HepG2 cells (Supplemental Figure 3, B and C), but NET marker SYTOX Green was absent in neutrophils that were simply cocultured with HepG2 cells (Supplemental Figure 3, D and E). These results suggest that NET formation is not responsible for ITPR2 degradation here. Since NETs are also a form of neutrophil death (42), neutrophils treated with methanol to cause cell death were cocultured with HepG2 cells, but this also did not reduce ITPR2 (Supplemental Figure 3, F–I). Together, these findings suggest that intact neutrophils must be in direct contact with HepG2 cells to decrease ITPR2 by a mechanism that is independent of integrins or NET formation.

Direct contact between HepG2 cells and intact neutrophils is important for the decrease in ITPR2. (A) Representative immunoblots and (B) quantitation of ITPR2 levels after 18–24 hours of culture of HepG2 cell in conditioned medium (C.M.) with neutrophils cultured for 16 hours. C.M. does not reduce ITPR2 in HepG2 cells. (C) Representative immunoblots and (D) quantitation of ITPR2 in HepG2 cells separated from neutrophils by a 3-μm pore Transwell system (T.W.) after 18–24 hours. Coculture using T.W. does not decrease ITPR2 in HepG2 cells. (E) Representative immunoblots and (F) quantitation of ITPR2 levels in HepG2 cells after 18–24 hours of coculture with naked neutrophils, from which cell surface GPI-anchored proteins were removed. Naked neutrophils are less effective in reducing ITPR2 in HepG2 cells. (G) Representative immunoblots and (H) quantitation of ITPR2 levels in HepG2 cells after preincubation with anti–integrin α2 (anti-ITGA2) or anti–integrin αM (anti-ITGAM) antibodies for 2 hours and coculture with neutrophils for 18–24 hours. Inhibition of these integrins does not alter the decrease in ITPR2 in HepG2 cells. (I) Representative immunoblots and (J) quantitation of ITPR2 levels after incubation of HepG2 cells with various concentrations of neutrophil extracellular trap (NET) components (neutrophil elastase, myeloperoxidase [MPO], high mobility group box-1 [HMGB1]) for 20 hours. Extracellular administration of these NET components does not alter ITPR2 in HepG2 cells. Data are presented as mean ± SD; n = 6 (F), n = 3–4 (J), n = 3 (B, D, and H). NS, not significant. *P < 0.05, **P < 0.01, ***P < 0.001 by 1-way ANOVA with Tukey’s multiple-comparison test.

Neutrophils insert granule proteins into hepatocytes. To investigate the neutrophil component that reduces ITPR2 in hepatocytes, cellular fractions of neutrophils were administered to HepG2 cells (Figure 4A) (43). ITPR2 was decreased by incubation with either the granule-containing fraction or the total cell homogenate, but not by the cytoplasmic or plasma membrane fractions of neutrophils (Figure 4, B and C). Moreover, the effect of total homogenate was eliminated by boiling (Figure 4, B and C). The same results were obtained using primary human hepatocytes and human neutrophils (Supplemental Figure 4, A and B). These findings suggest proteins within neutrophil granules are transferred to hepatocytes upon direct contact. To test this hypothesis, HepG2 cells were cocultured with neutrophils for 1 hour, followed by washing to remove neutrophils. Confocal immunofluorescence revealed that the neutrophil granule proteins MPO and neutrophil elastase were present and partially colocalized within the HepG2 cells (Figure 4, D and E). This was confirmed by immunoblotting (Figure 4, F and G). Transfer of MPO and elastase from neutrophils also occurred in human and mouse primary hepatocytes, as verified by immunoblotting and immunostaining (Supplemental Figure 4, C–J).

Neutrophils insert granule proteins into hepatocytes. (A) Flowchart showing the protocol for fractionating neutrophils. (B) Representative immunoblots and (C) quantitation of ITPR2 in HepG2 cells after 20 hours of incubation with the indicated fractions shows that the neutrophil granule fraction is sufficient to reduce ITPR2. (D and E) Representative confocal immunofluorescence images of HepG2 cells alone (D) and after coculture with neutrophils for 1 hour, after which the cells were then washed to remove the neutrophils (E). Labels are anti-myeloperoxidase (anti-MPO, green) and anti-elastase (red) antibodies, nuclear staining (DAPI, blue), and phalloidin (gray). Scale bar: 20 μm. Inset is an ×2 enlargement showing partial colocalization of MPO and elastase (yellow). Elastase also appears to be diffusely distributed in the nucleus. (F) Representative immunoblots and (G) quantitation show the neutrophil granule proteins MPO and elastase appear in HepG2 cells after coculturing with neutrophils for 1 hour, after which the cells were washed to remove the neutrophils. All data are presented as mean ± SD; n = 4–9 (C) and n = 3 (G). NS, not significant. ****P < 0.001 by 1-way ANOVA with Tukey’s multiple-comparison test.

Hepatocyte ITPR2 is degraded by neutrophil elastase and hepatocytes express Serpin E2 and A3 in response. In HepG2 cells cocultured with neutrophils, the decrease in ITPR2 was blocked by 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride (AEBSF), a serine protease inhibitor, but not by an MPO inhibitor (Figure 5, A–D). Administration of MPO to HepG2 cell homogenates rather than intact cells (Figure 3I) did not reduce ITPR2, but elastase at 0.2 μg/mL laddered the ITPR2 bands and eliminated them at 2.0 μg/mL (Figure 5, E and F). These results suggest neutrophil elastase is transferred to HepG2 cells by direct contact with neutrophils to degrade ITPR2. To examine the range of proteins degraded by neutrophil elastase (Figure 1K), proteomic analysis was performed using 2-dimensional difference gel electrophoresis (2D-DIGE) (44). Comparing HepG2 lysates with and without neutrophil elastase treatment, only 82 of 2153 spots were degraded by elastase (Figure 5G). Mass spectrometry identification of the 12 most degraded proteins is shown in Table 1. Infiltration of neutrophils occurs in most organs (22), so transfer of neutrophil granule proteins and ITPR2 degradation by elastase was examined in other cell types. In HCT116 (colon), A549 (lung), and PANC-1 (pancreas) cells cocultured with neutrophils for 1 hour, the presence of MPO and elastase was observed along with a decrease in ITPR2 (Supplemental Figure 5, A and B). Cathepsin G and MMP9, two other neutrophil granule proteins (45), could not be detected in HepG2 cells even after coculture with neutrophils (Supplemental Figure 5C). The observations that neutrophils do not damage hepatocytes in this system (Figure 1, A–E, and Supplemental Figure 1, B–K) and that removal of neutrophils restores ITPR2 (Figure 2, E–G) suggest hepatocytes have a mechanism to temporally limit this effect of neutrophils. To investigate this, bulk RNA-seq was performed using HepG2 cells 20 hours after incubation with either neutrophils or the neutrophil granule fraction (Supplemental Figure 5, D and E). We identified 101 upregulated and 108 downregulated genes by neutrophils, plus 50 upregulated and 49 downregulated genes by the granule fraction, respectively (Supplemental Figure 5F). Ten genes were downregulated by both neutrophils and granules, including Serpin E2 and Serpin A3 (Supplemental Figure 5F), 2 known antiprotease genes (46). Because proteolysis by neutrophil elastase in hepatocytes occurs as early as 1 hour after coculture, RT-qPCR was used to evaluate Serpin E2 and Serpin A3 gene expression in HepG2 cells at time points preceding 20 hours. Both mRNAs in HepG2 cells were significantly elevated at 1 and 4 hours of coculture with neutrophils (Figure 5, H and I), suggesting that HepG2 cells rapidly elevated Serpin E2 and Serpin A3 in response to neutrophil elastase, with a compensatory decrease in these mRNAs after 20 hours. Serpin A3 protein level also was elevated in human hepatocytes cocultured with neutrophils after 1 hour (Supplemental Figure 5, G and H). Finally, administration of recombinant Serpin E2 or Serpin A3 to HepG2 cell homogenates inhibited ITPR2 degradation by neutrophil elastase (Figure 5, J and K). These findings provide evidence that hepatocytes respond to contact with neutrophils by increasing expression of Serpin E2 and Serpin A3 to temporally limit the degradative effects of neutrophil elastase.

Hepatocyte ITPR2 is degraded by neutrophil elastase and hepatocytes express Serpin E2 and A3 in response. (A) Representative immunoblots and (B) quantitation of ITPR2 levels in HepG2 cells after coculture with neutrophils for 1 hour show that loss of ITPR2 is blocked by the serine protease inhibitor AEBSF (10 μM). (C) Representative immunoblots and (D) quantitation of ITPR2 levels in HepG2 cells after coculturing with neutrophils for 1 hour show that loss of ITPR2 is not blocked by the MPO inhibitor PF-1355 (10 μM). (E) Representative immunoblot and (F) quantitation of ITPR2 levels in HepG2 cell lysates 5 minutes after addition of MPO or neutrophil elastase at the indicated concentrations. Note the presence of multiple low molecular weight bands in homogenates treated with a low concentration of elastase, but ITPR2 is completely digested in homogenates treated with a higher concentration. (G) 2D-DIGE of a representative subproteome from lysates of HepG2 cells with (red, Cy5) or without (green, Cy2) neutrophil elastase (1 mg/mL). Yellow spots indicate overlap of both samples. Eighty-two spots were detected with a ratio of green to red fluorescence of greater than 1.5 (surrounded by circles), likely reflecting significantly degraded proteins. The 12 spots with the greatest differences (spots at the edges and spots near each other were avoided) were analyzed by mass spectrometry (Table 1). (H) RT-qPCR shows Serpin E2 mRNA is increased at early time points in HepG2 cells cocultured with neutrophils, compared with HepG2 cells alone. (I) RT-qPCR shows mRNA levels of Serpin A3 also are increased at early time points in HepG2 cells cocultured with neutrophils, compared with HepG2 cells alone. (J) Representative immunoblots and (K) quantitation of ITPR2 levels in HepG2 cell lysates 5 minutes after addition of neutrophil elastase protein (0.2 μg/mL) show that degradation of ITPR2 can be prevented by either recombinant Serpin E2 or Serpin A3 protein (20 μg/mL). All data are n = 3 and are presented as mean ± SD. NS, not significant. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001 by 1-way ANOVA with Tukey’s multiple-comparison test (B, D, F, and K) or unpaired, 2-tailed Student’s t test (H and I).

Protein identification by mass spectrometry and ratio of 12 spots (depicted in Figure 5G)

Granules containing MPO and elastase are taken up by hepatocytes by PI3K-mediated endocytosis. To investigate how neutrophils transfer granule proteins to HepG2 cells, immunoelectron microscopy was performed using gold nanoparticles conjugated with an anti-MPO antibody. Quantitative analysis of immunoelectron labeling showed that MPO in neutrophil granules migrates to HepG2 cells, where it was localized to an endosome-like structure (Figure 6, A and B). This endosome-like structure was in the cytoplasm of HepG2 cells cocultured with neutrophils, but not in control HepG2 cells (Supplemental Figure 6, A and B). Although this structure was larger than neutrophil granules, immunostaining showed that MPO partially colocalized with elastase, suggesting they migrate from neutrophils to hepatocytes in granules or vesicles (Figure 4E). Therefore, after removing cocultured neutrophils from HepG2 cells, the HepG2 cells were subjected to the same procedure that was used to isolate the granule fraction from neutrophils (Figure 4A). This resulted in recovery of a granule fraction from HepG2 cells that contained both MPO and elastase (Supplemental Figure 6C). Uptake of granule proteins and ITPR2 degradation could be inhibited by incubation at 4°C or treatment with PI3K inhibitors, LY294002 and wortmannin (Figure 6, C–F, and Supplemental Figure 6, D and E), suggesting granules are transferred by PI3K-mediated endocytosis. Colocalization of lysosome-associated membrane protein 1 (LAMP1) with neutrophil elastase in HepG2 cells cocultured with neutrophils (Figure 6, G and H) supports the notion that granule uptake is by the endocytosis pathway. Furthermore, in HepG2 cells after incubation of the neutrophil granule fraction with the lipophilic membrane dye PKH67 (47), the colocalization of elastase (Supplemental Figure 6, F and G) or LysoTracker (Supplemental Figure 6, H and I) with this lipophilic membrane dye also supports these results. Some of the neutrophil elastase in HepG2 cells did not colocalize with MPO (Figure 4E), appearing to diffuse into the nucleus (Figure 4E and Supplemental Figure 6G), which suggests the elastase is transported out of the internalizing granules. This phenomenon can occur when NETs are elicited in neutrophils and may be associated with MPO-induced reactive oxygen species (48). Administration of the antioxidant N-acetyl-L-cysteine (NAC) inhibited the degradation of ITPR2 (Supplemental Figure 6, J and K), which may suggest the possibility of a similar mechanism.

Granules containing MPO and elastase are taken up by hepatocytes via PI3K-mediated endocytosis. (A) Neutrophil granules are seen in HepG2 cells by transmission electron microscopy. Top panel: Representative image of HepG2 cells cocultured with neutrophils (N, nucleus; black asterisks, endosome-like structures in HepG2 cells). Scale bar: 1 μm. Bottom left panel: Magnified image shows gold nanoparticles (yellow arrows) bound to MPO antibodies in neutrophil granules. Scale bar: 500 nm. Bottom right panel: Magnified image shows MPO within endosome-like structures in HepG2 cells (yellow arrows). (B) Immunogold labeling is increased in HepG2 cells cocultured with neutrophils. Shown is the number of gold nanoparticles in HepG2 cells cultured alone or with neutrophils, or in neutrophils alone, each normalized by cell area. Data are from 14 images of HepG2 cells alone, 19 images of HepG2 cells cocultured with neutrophils, and 7 images of neutrophils alone. (C) Representative immunoblots and (D) quantitation of ITPR2 and MPO in HepG2 cells incubated with neutrophils or neutrophil granule fractions for 1 hour at 4°C or 37°C show that transfer of MPO and ITPR2 degradation are blocked at 4°C. (E) Representative immunoblots and (F) quantitation of ITPR2 and MPO in HepG2 cells incubated with LY294002 (40 μM) for 1 hour and then cocultured with neutrophils for 1 hour show that transfer of MPO and ITPR2 degradation are blocked by the PI3K inhibitor. (G) Representative confocal images of HepG2 cells alone and (H) after coculture with neutrophils for 1 hour. Labels are anti-LAMP1 (green) and anti–neutrophil elastase (red) antibodies, Hoechst 3342 (blue), and phalloidin (gray). Scale bar: 20 μm. Note that in cocultured HepG2 cells, some LAMP1 colocalizes with neutrophil elastase (yellow; better appreciated in ×2 magnification in inset). All data are presented as mean ± SD; n = 3 (D and F). NS, not significant. **P < 0.01; ***P < 0.001; ****P < 0.0001 by unpaired, 2-tailed Student’s t test (B) or 1-way ANOVA with Tukey’s multiple-comparison test (D and F).

Loss of ITPR2 and impaired hepatocyte proliferation are attenuated in an elastase-deficient mouse model of AAH. Ethanol-LPS-fructose (ELF) treatment was used as a mouse model of AAH (37) and it showed that ITPR2 levels in liver homogenates were significantly reduced compared with controls (Supplemental Figure 7, A and B). The number of neutrophils infiltrating the liver was significantly increased as well (Supplemental Figure 7, C–E). Serpin E2 mRNA of ELF-treated liver also increased compared with controls (Supplemental Figure 7F). The chronic-plus-binge AAH model (49) also resulted in decreased liver ITPR2 levels compared with controls (Supplemental Figure 7, G and H). There was also an increase in infiltrating neutrophils over controls in this model, but it was less pronounced than in the ELF model (Supplemental Figure 7I). These results indicate there is neutrophilic infiltration and decreased ITPR2 in the livers of AAH model mice. To clarify the relationship between neutrophil elastase and ITPR2 degradation in vivo, protein levels of primary mouse hepatocytes were compared after 1 hour of coculture with neutrophils isolated from neutrophil elastase–knockout (Elane–/–) or wild-type (WT) mice. MPO was found in both hepatocytes, but Elane–/– neutrophils failed to degrade ITPR2 (Figure 7, A and B). Next, the ELF model was examined in Elane–/– and WT mice (Figure 7C). ITPR2 levels in the livers of Elane–/– mice were significantly increased compared with WT, as was the hepatocyte proliferation marker cyclin D1 (Figure 7, D and E) (26). The number of hepatocytes expressing Ki67, another cell proliferation marker, was also significantly higher in Elane–/– than WT mice (Figure 7, F–H). In WT ELF-treated mice, treatment with the neutrophil elastase–specific inhibitor sivelestat or with AEBSF also increased the number of Ki67-positive hepatocytes (Supplemental Figure 7, J–M), and the levels of ITPR2 and cyclin D1 in the liver also were significantly increased (Supplemental Figure 7, N–O). Finally, serum ALT levels were higher in ITPR2–/– ELF mice than in matched WT ELF mice (145 ± 42 vs. 101 ± 26 U/L, P = 0.02, one-tailed Student’s t test) and lower in Elane–/– ELF mice than in matched WT ELF mice (56 ± 8 vs. 81 ± 24 U/L, P = 0.006, one-tailed Student’s t test). Collectively, these results provide evidence that ITPR2 in hepatocytes is degraded, and proliferation is reduced by elastase from infiltrating neutrophils in the ELF model of AAH. Finally, the livers of ITPR2–/– ELF mice had significantly lower levels of cyclin D1 and significantly fewer Ki67-positive hepatocytes compared with WT ELF mice (Figure 7, I–K). This is consistent with previous reports that decreased hepatocyte proliferation is related to ITPR2 deficiency (18) and extends this to AAH.

Loss of ITPR2 and impaired hepatocyte proliferation are attenuated in an elastase-deficient mouse model of alcohol-associated hepatitis (AAH). (A) Representative immunoblots and (B) quantitation of ITPR2, MPO, and elastase in primary mouse hepatocytes after 1 hour of coculture with neutrophils from WT or neutrophil elastase–KO (Elane–/–) mice. Findings support the notion that granules are transferred to hepatocytes from either type of neutrophil, but ITPR2 is only reduced if elastase is present. (C) Representative immunoblots confirm expression of MPO but not elastase in bone marrow of mice transplanted with Elane–/– or WT bone marrow, respectively. (D–H) In AAH models, ITPR2 levels, which are reduced in WT mice, improve in Elane–/– mice and hepatocyte proliferation increases. AAH was induced by the ethanol + LPS + fructose (ELF model; ref. 41) in Elane–/– or WT mice. (D) Representative immunoblots and (E) quantitation of ITPR2 and cyclin D1 in liver homogenates show that both are increased in Elane–/– mice. The lanes were on the same gel but noncontiguous. Representative confocal images of (F) WT and (G) Elane–/– livers immunostained with anti-CK18 (green) and anti-Ki67 (red) antibodies. Scale bar: 50 μm. (H) Quantitative comparison confirms the number of Ki67-positive hepatocytes is greater in Elane–/– mice. (I–K) In the AAH (ELF) model, hepatocyte proliferation is decreased in ITPR2-deficient (ITPR2–/–) mice compared with WT mice. (I) Representative immunoblots and (J) quantitation of the levels of ITPR2 and cyclin D1 in liver homogenates show a reduction of cyclin D1 in ITPR2–/– mice. (K) Quantitation of immunostaining of livers with anti-CK18 and anti-Ki67 antibodies shows that the number of Ki67-positive hepatocytes is reduced in the KO mice. All data are expressed as mean ± SD; n = 4 (B), n = 8 (E), 4 fields of view per mouse (H), n = 7 for WT and n = 5 for ITPR2–/– (J), and 4–5 fields of view per mouse (K). NS, not significant. **P < 0.01; ***P < 0.001; ****P < 0.0001 by 1-way ANOVA with Tukey’s multiple-comparison test (B) or unpaired, 2-tailed Student’s t test (E, H, J, and K).

Degradation of ITPR2 in hepatocytes occurs by insertion of neutrophil elastase in patients with AAH. To determine the relevance of these observations to human disease, liver biopsies from patient specimens that were histologically normal were compared to specimens from patients with biopsy-proven AAH. Immunohistochemical staining for neutrophil elastase and MPO showed a significant increase in the number of positive cells found in the liver parenchyma from patients with AAH compared with histologically normal controls (Supplemental Figure 8, A–F), while immunohistochemical staining for ITPR2 was significantly decreased in hepatocytes of biopsies from patients with AAH, relative to histologically normal controls (Figure 8, A–C). Protein levels were evaluated in homogenates of liver explants from patients transplanted for severe AAH, plus histologically normal livers (patient information in Table 2). Consistent with in vitro findings, ITPR2 and SERCA2 were significantly decreased and MPO and elastase were significantly increased in AAH compared with normal livers (Figure 8, D and E). Moreover, ITPR2 staining was inversely related to the number of elastase-positive cells (r = –0.63, P = 0.03; Figure 8F). Furthermore, higher magnification identified MPO- or elastase-positive granular staining in hepatocytes near neutrophils (Supplemental Figure 8, G and H), which was more apparent with fluorescent immunostaining for CK18 (hepatocytes), neutrophil elastase, and MPO in normal or AAH liver sections (Figure 8, G and H). In histologically normal specimens, neutrophil elastase and MPO were mostly confined to sinusoidal regions, whereas in AAH specimens, they appeared as punctate labels within CK18-positive cytoplasm, some of which colocalized. The number of elastase-positive puncta inside hepatocytes was significantly higher in AAH specimens than in controls (Figure 8I). Serpin A3 levels were significantly higher in homogenates of AAH liver samples than in controls (Supplemental Figure 8, I and J) as well. Finally, AAH specimens were fluorescently labeled for CK18, neutrophil elastase, and the apoptosis marker cleaved caspase 3 to detect perforocytosis (50). The occurrence of neutrophils in apoptotic hepatocytes was observed (Supplemental Figure 8K), but was rare.

Degradation of ITPR2 in hepatocytes occurs by insertion of neutrophil elastase in patients with alcohol-associated hepatitis (AAH). (A–C) ITPR2 is reduced in hepatocytes in patients with AAH. Representative images of immunohistochemical staining with anti-ITPR2 antibody in (A) histologically normal controls (Ctrl) and (B) liver biopsy specimens from patients with AAH. Inset in each image is magnified ×3. Scale bar: 50 μm. (C) Quantitative measurement of ITPR2 staining confirms decreased expression in AAH. Three fields were quantified in each biopsy specimen (8 Ctrl and 9 AAH). (D–F) There is an inverse relationship between ITPR2 in hepatocytes and neutrophil elastase in liver biopsies from patients with AAH. (D) Representative immunoblots and (E) quantitation of ITPR2, SERCA2, MPO, and elastase in homogenates of livers from 8 Ctrl and explants from 8 patients who received liver transplants for AAH. ITPR2 and SERCA2 are decreased further in patients with more neutrophil infiltration. (F) There is an inverse relationship between ITPR2 and neutrophil elastase staining (r = –0.63, Spearman’s correlation; P < 0.05) based on the 12 individuals who underwent both analysis of the immunostained region of ITPR2 (A–C) and the number of elastase-stained positive neutrophils in panel (D–F) of Supplemental Figure 8. (G–I) Neutrophil elastase is found in hepatocytes of patients with AAH. Representative images of confocal fluorescent immunostaining with anti-CK18 (gray), anti-MPO (green), and anti-elastase (red) antibodies from (G) histologically normal human liver and (H) liver of AAH patient. Yellow asterisks represent neutrophils and arrowheads indicate that MPO and elastase dots are present within CK18-positive hepatocytes. Inset in H is an ×2 magnified image showing partial colocalization of MPO and elastase (yellow). Scale bar: 20 μm. (I) The number of spots of elastase present inside CK18-positive hepatocyte regions is significantly increased in AAH biopsies relative to controls. Ctrl = 8, AAH = 8. All data are presented as mean ± SD. *P < 0.05, **P < 0.01, ***P < 0.001 by 1-tailed Student’s t test (C) or unpaired, 2-tailed Student’s t test (E, F, and I).

Clinical information for patients with AAH whose frozen liver sections were analyzed

Neutrophils from patients with AAH are more potent than healthy controls in degrading ITPR2. The function of neutrophils in patients with AAH may differ from that of healthy donors (51, 52), and indeed we found that neutrophils from AAH patients (information in Table 3) were more capable than neutrophils from healthy individuals in degrading ITPR2 in HepG2 cells (Figure 9, A–D). MPO and elastase levels were similar in healthy and AAH neutrophils, suggesting that differences between the neutrophils are not due to increased expression of granule proteins (Figure 9, E–G). Therefore, ERK phosphorylation in response to stimulation with N-formyl-methionyl-leucyl-phenylalanine (fMLP) was evaluated as an index of neutrophil activation, including degranulation (53). Indeed, AAH neutrophils displayed increased ERK phosphorylation when compared with control neutrophils (Figure 9, H and I). These results suggest that neutrophils in AAH are primed (54) to transfer granule contents into hepatocytes, resulting in a more effective degradation of ITPR2.

Neutrophils from patients with alcohol-associated hepatitis (AAH) are more capable than healthy controls of degrading ITPR2. (A–D) Kinetics of ITPR2 loss in HepG2 cells cocultured with control or AAH neutrophils. (A) Representative immunoblots of ITPR2 levels in HepG2 cells cocultured with neutrophils for 20 hours from (A) healthy volunteers or (B) patients with AAH. (C) Quantification of data from 7 healthy volunteers and 7 AAH patients shows a significant difference in measured ITPR2/GAPDH levels when 0.025 × 106 or 0.05 × 106 neutrophils were administered. (D) First-order rate constant K is higher for AAH neutrophils than for controls. (E–G) MPO and elastase content is similar in control and AAH neutrophils. (E) Representative immunoblots and (F and G) quantitation of MPO and elastase protein levels in neutrophils from 7 healthy volunteers and 7 AAH patients. (H and I) ERK signaling is more active in AAH neutrophils. (H) Representative immunoblots and (I) quantitation of p-ERK and ERK after stimulation of neutrophils isolated from blood of 8 healthy volunteers (Ctrl) or 7 AAH patients with fMLP (100 nM) for 5 minutes. Data are shown as mean ± SEM (C and D) or mean ± SD (F, G, and I). NS, not significant. *P < 0.05, **P < 0.01 by unpaired, 2-tailed Student’s t test.

Clinical information for patients with AAH whose neutrophils were analyzed