Induction of Vesicular Trafficking and JNK-Mediated Apoptotic Signaling in Mononuclear Leukocytes Marks the Immuno-Proteostasis Response to Uremic Proteins

Chronic kidney disease (CKD) is a condition of solute retention [1]. Conventionally classified as small, medium, and large sizes [2], these solutes sustain uremic symptoms, including immune dysfunction and its clinical correlates. These are main complications of CKD and especially of end-stage renal disease, leading to defective response to the host, increased risk of auto-immune diseases, altered response to vaccines, and poor control of inflammatory processes resulting in low-grade inflammation and vascular damage. Leukopenia is a characteristic cue of immune dysfunction in CKD, that mainly presents as a moderate to severe lymphocytopenia, by the premature death of peripheral blood mononuclear leukocytes (PBL) essentially by apoptosis [3‒8]. However, such uremic complication involves all the leukocyte subsets, including lymphocytes, monocytes, and polymorphonuclears [9, 10], and increased levels of apoptosis are also observed in bone marrow progenitors of the myeloid lineage and even in peripheral red blood cells [11‒13], eventually sustaining both uremic leukopenia and anemia [14]. An increased cell death has also been reported in other tissues, such as arterial endothelium ([15] and references therein) and bones (reviewed in [16]), suggesting that this may represent a systemic and disease-specific process of tissue degeneration and loss of cell mass.

Earliest studies carried out in hemodialysis (HD) patients treated with protein-leaking dialyzers (PLD) suggested that large (proteinaceous) solutes could play a main role in PBL death [17]. More recently, protein-bound uremic toxins, i.e., small solutes that covalently bind to plasma proteins, such as p-cresol and indoxyl sulfate, have been described to behave as pro-apoptotic agents [15, 18‒20]. Other large or high-molecular-weight solutes (HMW) that might intervene in the premature death of PBL in CKD patients include inflammatory mediators [21, 22]. Fas ligand, a component of the TNF family of membrane-bound proteins, is a pro-apoptotic cytokine found in uremic serum [23], and pro-apoptotic effects have also been demonstrated for other cytokines involved in the low-grade inflammatory syndrome of CKD [6, 24].

Again, cytotoxic and pro-inflammatory solutes that might promote cell death in uremic tissues are formed by the reduced renal clearance and metabolism of advanced oxidation and glycation products of plasma proteins [25, 26] as well as of advanced oxidation protein products and protein carbonyls [27‒29]. These protein decoration products coexist in uremic plasma (u-Pl), leading to a severe alteration of protein homeostasis (proteostasis) and activation of scavenging processes by the reticuloendothelial system, the vascular endothelium, and other cellular components of different tissues (reviewed in [29‒31]). Receptor-dependent and independent mechanisms of endocytosis are involved in these scavenging processes that characterize human diseases associated with a defective proteostasis. Receptor-dependent mechanisms include the activation of the receptor for advanced glycation end-products (RAGE), a nonspecific multi-ligand pattern recognition receptor widely investigated in diabetes and uremia for its role in low-grade inflammation and endothelial damage (reviewed in [32, 33]). In the present study, the role of uremic HMW (u-HMW) as instigators of premature death in uremic PBL (u-PBL) was investigated ex vivo in PBL and in vitro in mononuclear cell lines, starting from the hypothesis that an increased scavenging of large uremic solutes may induce apoptosis interfering with stress response and proteostasis mechanisms of these cells.

Ten CKD patients with end-stage renal disease enrolled at the HD unit of “Fondazione IRCCS Policlinico San Matteo” in Pavia, Italy, were included in this proof-of-concept study. Exclusion criteria included recent illness (within the previous 2 months); significant anemia (Hb < 10 g/dL); autoimmune disease; but also systemic diseases as vasculitis, amyloidosis, rheumatic disease; HBV, HCV, HIV positivity or other active viral positivity; active bacterial infection; active or previous cancer remission; previous transplantation.

CKD patients were on HD, 4 h thrice weekly, with non-reused standard low-flux polysulfone dialyzers. The dialytic procedure was standard bicarbonate dialysis, and the Kt/V index ranged from 1.24 to 1.35. The surface of the filters was tailored to the patients’ needs, and the patients were clinically stable; more in detail, no relevant changes occurred in the plasma levels of urea, phosphate, calcium, and creatinine, or in hematocrit, Kt/V, and leukocyte counts. No patient was under treatment with drugs acting directly on the immune system, such as steroids or other immunosuppressants. Blood samples were collected from patients just before starting the dialysis session at the end of the long interdialytic interval. At inclusion, patients presented with normal WBC count (6.9 ± 1.8 cells 10³/μL; reference value 3.6–9.6 cells 10³/μL) and mild lymphopenia (median ± SD and range of lymphocyte counts were 1.3 ± 0.6 and 0.3–2.9 cells 10³/μL, respectively, and median ± SD and range of lympocyte % were 18.3 ± 7.1 and 7.7–29.8, respectively; corresponding values in healthy controls of lymphocyte % were 36.6 ± 6.8 and 20.5–51.1).

Teenage and sex-matched hospital staff and healthy individuals were also included in the study as controls. Participant characteristics are summarized in Table 1.

Blood was collected from CKD patients and healthy control subjects in Vacutainer^® tubes containing sodium heparin as anticoagulant, and PBL were isolated by density gradient centrifugation using Lympholyte-H (Cedarlane, Burlington, ON, Canada) according to the manufacturer’s protocol. Briefly, whole blood was diluted 1:1 in sterile saline, layered onto Lympholyte-H cell separation medium, and centrifuged at 870 g for 30 min at room temperature without brakes. The layer containing PBL between the upper (plasma, thrombocytes) and the lower phase (Lympholyte-H) was carefully collected using a Pasteur pipette, transferred to a fresh tube, and washed twice with phosphate-buffered saline (PBS). The PBL pellets were suspended in ammonium chloride-potassium lysing buffer (Invitrogen; Thermo Fisher Scientific, Milan, Italy) and incubated for 10 min at room temperature with gentle mixing to lyse contaminating RBC. This was followed by a wash of the purified cells with PBS-EDTA.

In some experiments, the effects of u-Pl and its HMW fraction, prepared as described later, were investigated in control PBL (c-PBL) obtained from the local blood bank. Outdated blood bags containing buffy coat intended for disposal were obtained on the day of the expiry date and treated for PBL isolation using the same Lympholyte-H density gradient isolation procedure described earlier for fresh PBL.

u-PBL and c-PBL were maintained at 37°C in a humidified atmosphere containing 5% CO₂ suspended in RPMI-1640 (GIBCO, Life Technologies) containing 1% l-glutamine and 10% vol/vol fetal bovine serum (FBS, GIBCO, Life Technologies). For the treatments, u-Pl or healthy control plasma (c-Pl) or their HMW fractions were used at 10% vol/vol instead of FBS. A hemocytometer was used to count the total cell number, and cell viability was assessed by Trypan blue exclusion test using a solution of 0.4% w/vol of this dye that was obtained from Sigma-Aldrich, St. Louis, MO, USA.

To study the effects of uremic retention solutes on PBL viability, a fraction enriched of these solutes was prepared starting from u-Pl and c-Pl by in vitro ultrafiltration (u-HMW and c-HMW, respectively). Micro-concentrator membranes (Vivaspin 6 mL, Sartorius Stedim Biotech GmbH, Germany) with 30 and 50 kDa nominal cutoff were used for this solute enrichment procedure according to manufacturer’s instructions. The 50-kDa MW cutoff is expected to contain the majority of large solutes with cell death-inducing activity in u-Pl since dialyzer membranes used in both diffusive and convective HD methods and in vitro ultrafiltration procedures based on membrane microconcentrators with nominal cutoff ranging between 10 and 30 kDa unvariably showed limited efficacy in removing them [4, 34]. At the same time, preliminary data suggested that plasma protein fractions enriched of large solutes prepared by ultrafiltration carried out either in vivo in patients treated with protein-leaking (PL) dialyzers (nominal cutoff >50 kDa) or in vitro using 50 kDa cutoff micro-concentrators, contain most of the proteinaceous solutes that cause DNA damage and cell death in mononuclear cell lines [35]. To further confirm these findings, both the 50 kDa and 30 kDa cutoff fractions (u-HMW^50kDa and u-HMW^30kDa, respectively) were compared with whole plasma for their effects on PBL viability and other cell and molecular parameters investigated in this study.

Protein carbonyls were investigated by immunoblot and spectrophotometric methods as described ([36] and references therein) using the Brady’s reagent 2,4-dinitrophenylhydrazine (2,4-DNPH), according to Levine et al. [37]. More in detail, the immunoblot analysis of 2,4-DNPH-reactive carbonyls was performed in plasma proteins separated by two-dimension electrophoresis (2D-PAGE) performed under denaturation conditions and transferred by electroblotting onto a nitrocellulose membrane for identification after derivatization using an anti-2,4-DNPH antibody (Abcam, Cambridge, UK). For the first dimension of 2D gradient PAGE analyses, from 100 to 300 mg of total proteins were loaded onto immobilized pH gradient gel strips (17 cm, pH 3–10 NL), and isoelectric focusing was performed using a Protean IEF cell system (Bio-Rad, Hercules, CA, USA). The second dimension was performed on 9–16% polyacrylamide linear gradient electrophoresis gels using a Protean II xi 2-D System (Bio-Rad). The gels were stained with silver stain, and image analysis was performed using the PD-Quest 2D gel software (Bio-Rad, Hercules, CA, USA). Reference 2D maps of human plasma proteins were obtained from SWISS-PROT database (https://world-2dpage.expasy.org/swiss-2dpage) and protein identity was confirmed by LC-MS analysis after protein isolation by manual spot picking and in-gel digestion using MS grade trypsin as described in [26, 38].

THP-1 human monocytic leukemia cells and human K562 erythroleukemic cell line (American Type Culture Collection, ATCC, Manassas, VA, USA) maintained in RPMI, as described earlier for human PBL, were used at intervals of passage in culture between 10 and 25. For macrophage differentiation, THP-1 cells seeded in 6-well plates (10⁶ cells/mL) were treated with 5 ng phorbol-12-myristate-13-acetate (PMA)/mL for 48 h. After differentiation, the cells were washed 3 times with PBS and used for the incubation with plasma samples and their HMW fractions. These were added to the cell culture media of mononuclear cell lines and differentiated macrophages instead of FBS that served as negative control treatment.

Peripheral blood mononuclear leukocytes from healthy controls or CKD patients were seeded (15,000 cells/well) for the incubation (2 h) with autologous or heterologous plasma or HMW fractions and then the cells were stained with 0.4% trypan blue dye to assess in triplicate viable and nonviable cells. Cell viability of THP1 and K562 cells was assessed by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT, Sigma-Aldrich, St. Louis, MO, USA) assay performed according to the procedure previously published in [39] or using a Via1-Cassette™ (ChemoMetec) kit with NucleoCounter^® NC-3000™. The cassettes were loaded with immobilized acridine orange (AO) and DAPI for labeling of total cell and dead cell numbers, respectively.

Apoptosis was assessed by cytofluorimetric analysis using an Attune NxT Acoustic Focusing Cytometer (Thermo Fisher Scientific); after incubation for 24 h with plasma samples or their HMW fractions, THP1 cells (2 × 10⁵/well) were harvested and stained with Annexin V, Alexa Fluor™ 488 conjugate (Thermo Fisher Scientific), and propidium iodide (PI, Clontech), and then apoptosis was determined according to the manufacturers’ instructions. Staurosporine (50 μm, Sigma-Adrich, St. Louis, MO, USA) was used for positive control tests.

Autophagy was studied in THP1 cells by flow cytometry analysis (Attune NxT Acousting Focusing Cytometer; Thermo Fisher Scientific) and microplate immunofluorescence microscopy (Operetta CLS, Perkin Elmer). Autophagic vacuoles were identified utilizing a Cyto-ID Autophagy Green Detection Kit (Enzo Life Sciences, Inc.) and rapamycin and overnight starvation (cell culture in the absence of FBS) were used as positive controls.

THP-1 cells were grown in 6-well plates and treated with whole plasma or HMW fractions. The cells (1 × 10⁶/well) were suspended in 200 μL PBS, fixed by adding 800 μL of iced 100% ethanol, and then incubated overnight at 4°C. The cell pellets were collected by centrifugation and resuspended in hypotonic buffer containing 5 μg/mL RNase and 50 μg/mL propidium iodide and incubated for 1 h at 37°C. Fluorescence emitted from the propidium iodide-DNA complex was quantified after excitation of the fluorescent dye by NucleoCounter^® NC-3000™.

PBL, THP-1 cells, and THP-1 macrophages were treated with plasma or HMW fractions for 3 h and then washed twice before incubation in 50 μm DCFH-DA solution (Sigma-Aldrich, St. Louis, Missouri, USA) for 30 min at 37°C in the dark. Then the cells were washed with PBS, and the fluorescence was recorded using a DTX880 Multimode Detector microplate reader (Beckman Coulter).

Extracellular reactive oxygen species (ROS) were determined using the Amplex™ Red Hydrogen Peroxide/Peroxidase Assay Kit (Invitrogen). Briefly, 100 μL of cell culture medium were collected in 96-well plates for the incubation (10 min, at 37°C) in a humidified atmosphere with 5% CO₂ in the presence of 25 μL of HRP (1 U/mL) and 25 μL of Amplex red reagent. Then, the fluorescence was measured at λ_excitation = 560 ± 20 nm and λ_emission = 585 ± 20 nm in a DTX880 Multimode Detector microplate reader (Beckman Coulter). The assay was calibrated with authentic H₂O₂.

Cells were resuspended in 200 μL of ice-cold cell lysis buffer containing 20 mm Tris-HCl (pH 7.5), 150 mm NaCl, 1 mm Na₂EDTA, 1 mm ethylene glycol-tetra-acetic acid, 1% Triton, 2.5 mm sodium pyrophosphate, 1 mm β-glycerophosphate, 1 mm Na₃VO₄, and 1 μg/mL leupeptin (Cell Signaling Technology), and 20 μL/mL protease inhibitor cocktail (Pierce, Thermo Fisher Scientific). After centrifugation at 14,000 g for 20 min at 4°C, the supernatant containing cellular proteins was collected and the BCA protein assay (Pierce, Thermo Fisher Scientific) was used to measure protein concentrations; bovine serum albumin was used as an external calibration standard.

After extraction, the protein samples were first resolved by 4–12% SDS-PAGE and then transferred to nitrocellulose membrane for immunoblot analysis as described in [40]. Primary antibodies from Cell Signaling Technology utilized in this study were anti-GAPDH (#5174) and β-actin (#3700) that were used to normalize immunoblot data, and anti-c-Jun (#9165), anti-phospho-ERK1/2 (#4377), anti-ERK1/2 (#4695), anti-phospho-SAPK/JNK (#4671), anti-JNK (#9258), anti-PERK (C33E10) (#3192; 1:1,000), anti-IRE1α (#3294, 1:1,000). Anti-transferrin (YIF-LF_MA0242; 1:500), anti-fibrinogen (YIF-LF_MA0108; 1:2,000), and anti-human serum albumin (1:1,000) were from AdipoGen Life Sciences.

Peroxidase-conjugated secondary antibodies were anti-rabbit (#7074) or anti-mouse (#7076) IgG HRP-linked antibody (Cell Signaling Technology). The signals were visualized by enhanced chemiluminescence reagent (ECL Plus, Pierce, or ECL Clarity Max, Bio-Rad), and band quantification was performed by Gel-Pro Analyzer.

Data were presented as mean ± SD, unless otherwise specified; study variables were processed for analysis of variance by ANOVA test, and differences between mean values in the experimental groups were assessed using Student’s t test (two-sided test for normally distributed data) or Mann-Whitney test (for nonparametric data), when appropriate; p values <0.05 were accepted. Correlations between continuous dependent variables were assessed using the Pearson’s correlation test.

• 1.

  Proteomics studies and carbonylation pattern of HMW fraction

A representative 2D-PAGE protein pattern of u-Pl is shown in Figure 1a. The identity of the main protein species was determined by image analysis using reference 2D-PAGE images of plasma proteins available at SWISS-PROT and internal proteomics database; the latter includes images of u-Pl and its HMW fraction rich in posttranslational modifications (PTM) prepared by in vivo ultrafiltration in CKD patients treated with PLD [26, 29, 38, 41].

Protein carbonylation was investigated by immunoblot (Fig. 1b) and spectrophotometric analyses (Fig. 1c) in u-Pl and c-Pl after derivatization with 2,4-DNP. Protein carbonyls were used as a reporter epitope to epitomize the multiparameter decoration process of large proteins in u-Pl [27, 29]. Since this and other processes of protein decoration have been alleged to sustain leukocyte activation and damage by different, and so far poorly understood, mechanisms (presented in the Introduction section and reviewed in [26, 31]), in this study we explored the association between protein decoration and the pro-apoptotic activity (presented later) of large solutes contained in u-HMW^50kDa. Average total levels of 2,4-DNP-reactive material in u-Pl measured by spectrophotometric analysis were several folds those observed in c-Pl, confirming a condition of defective proteostasis in CKD patients (Fig. 1c). At the LC-MS analysis, the main 2,4-DNP-reactive plasma proteins identified by 2D immunoblot (Fig. 1b) were confirmed to include albumin (primary accession number: P02768), transferrin (serotransferrin or siderophilin, P02787), and the fibrinogen chain α (P02671) and β (P02675). Less abundant carbonylated species were haptoglobin (alpha and beta chains, P00738), angiotensinogen (serpin A8; P01019), and antithrombin-III (SERPIN C1; P01008), α1-antitrypsin (also known as serum trypsin inhibitor or SERPIN A1; P01009), immunoglobulin heavy chain μ (intermediate segment M, P99009), and the secretory IgG chain A (P99003).

• 2.

  u-HMW solutes reduce cell viability and induce apoptosis in u-PBL

Freshly isolated uPBL maintained in FBS showed a trend toward a reduced cell viability compared to c-PBL (Fig. 2a). These freshly isolated PBL were utilized to assess ex vivo the effect of uPl and u-HMW on cell viability. In these experiments, u-Pl significantly reduced cell viability compared to c-Pl in the PBL of both the groups; treatments with u-HMW^50kDa almost completely reproduced the effect of u-Pl on c-PBL viability and further aggravated the effect of u-Pl on the viability of u-PBL (Fig. 2a, left chart and right chart, respectively).

These results demonstrate an increased susceptibility of uremic leukocytes to large uremic solutes with cell death-promoting activity which could mirror a priming effect by these solutes occurring in vivo for the u-PBL or a lower capability of these cells to adapt to uremic stressors by the allostatic load of uremia and dialysis adverse effects. Important was the observation that cell treatments with u-HMW^30kDa (online suppl. Fig. S1A, right chart, and online suppl. Fig. S1; for all online suppl. material see https://doi.org/10.1159/000533309) have much lower impact on cell viability compared to u-HMW^50kDa, which confirms that the large solutes of u-Pl responsible for the reduced cell viability of PBL mainly present a MW >50 kDa [17]. When investigated by flow cytofluorometry, the cell viability reduction effect of u-Pl in PBL was found to depend on increased levels of apoptosis; this mainly presented as late apoptosis (Fig. 2b, c), and solutes present in the u-HMW^50kDa fraction enhanced the pro-apoptotic effect of u-Pl (Fig. 2d).

• 3.

  Pro-apoptotic activity of u-HMW in mononuclear cell lines

THP1 monocytes and K562 erythroleukemia cells were used as mononuclear cell lines to further explore the pro-apoptotic activity of u-Pl and its u-HMW^50kDa fraction (Fig. 3). u-Pl and u-HMW^50kDa significantly reduced cell viability in both the cell lines (Fig. 3a, d), recapitulating the findings on PBL viability presented earlier in this section. The superiority of u-HMW^50kDa with respect to u-HMW^30kDa in producing this adverse effect on cell viability was also confirmed in these cell lines (online suppl Fig. S1C).

Cytofluorimetric data demonstrated that the effect of u-HMW^50kDa on THP-1 cell viability depends on the induction of both apoptosis and necrosis (Fig. 3b), and increased levels of DNA fragmentation were observed upon exposure to such uremic solutes in both THP1 (Fig. 3c) and K562 cell lines (Fig. 3f) indicating the entrance of these mononuclear cells in the execution phase of the apoptotic program, i.e. the sub-G1 phase of the cell cycle. u-HMW rapidly activated cell death mechanisms of mononuclear leukocytes, with average reduction of cell viability in K562 cells of approx. 70% after 3 h of incubation (Fig. 3e). Furthermore, confocal microscopy and flow cytofluory data demonstrated that u-HMW^50kDa induces higher levels of autophagy in THP-1 cells compared to c-HMW^50kDa, and a trend toward increased levels of autophagy was also observed in the comparison between u-HMW^50kDa and u-Pl (Fig. 4).

• 4.

  u-HMW stimulates H₂O₂ production and efflux in PBL and monocyte-macrophage cell lines

The levels of cellular ROS in mononuclear leukocytes can vary in response to several physiological and pathological stimuli, reflecting an increased activity of cellular oxidoreductases and/or the induction of cellular damage and oxygen activation in organelles such as mitochondria, peroxisomes, and the ER. Changes of the cellular redox induced by these changes in the flux of cellular ROS may lead to oxidative stress and activation of apoptosis [42] and/or other cell death programs [43].

Compared to c-Pl, the treatment with u-Pl significantly increased ROS levels in c-PBL, and these levels further increased when the cells were treated with u-HMW^50kDa (Fig. 5a), but not with u-HMW^30kDa (online suppl. Fig. S1D). This oxygen activation response to u-Pl and u-HMW^50kDa treatments was recapitulated in THP1 monocytes (Fig. 5b) and K562 cells (online suppl. Fig. S2).

To better characterize the nature of these cellular ROS, c-PBL and THP-1 monocytes were loaded with the H₂O₂-scavenging enzyme catalase and then treated with u-HMW^50kDa. These experiments demonstrated that the cellular ROS formed during u-HMW^50kDa treatment mainly consist of H₂O₂ (Fig. 5a, left panel). Also, we found that this ROS once formed is secreted out of the cell (Fig. 5a, right panel). These findings were confirmed in THP-1 cells, also demonstrating that the induction effect of u-HMW^50kDa on the cellular synthesis and efflux of H₂O₂ was enhanced in this cell line by macrophage differentiation (Fig. 5c, right panel).

• 5.

  u-HMW solutes accumulate in u-PBL and activate the UPR/ER stress response (ERSR) of these cells

Uremic toxins, such as indoxyl sulfate, p-cresyl sulfate, and indole acetic acid, have recently been found to accumulate in endothelial cells, also promoting cellular accumulation of protein carbonyls and a defective autophagy [20]. To investigate whether u-HMW^50kDa induce the same effects in PBL, the main proteins and carbonylation targets of this HMW fraction, i.e., transferrin, albumin, and fibrinogen (Fig. 1), were searched by immunoblot in the total protein extract of u-PBL. The results demonstrated the presence of these proteins in u-PBL, but not in c-PBL (Fig. 6a), which can be explained by an increased cellular scavenging of these uremic proteins.

This cellular accumulation of plasma proteins was associated with increased levels of expression of the UPR and ER stress-signaling proteins IRE1-α and PERK (Fig. 6b), as well as of SAPK-JNK phosphorylation and c-Jun expression (Fig. 6c). At the same time, MAPK-ERK1/2 phosphorylation showed a trend toward a reduction (Fig. 6d). u-HMW^50kDa treatment in THP-1 cells recapitulated the SAPK-JNK and MAPK-ERK1/2 phosphorylation patterns of u-PBL (online suppl. Fig. S3).

Uremic retention solutes have long been described to play a causal role in uremic leukopenia and immune dysfunction by activating the apoptotic program of both polymorphonuclear and mononuclear leukocytes [9, 10]. Such a pro-apoptotic activity has also been described in other cell types, such as bone cells (reviewed in [16]), vascular endothelial cells, and resident phagocytes of uremic tissues (reviewed in [15, 20] and references therein).

In an attempt to better characterize the nature and the mechanism of action of these pro-apoptotic solutes, we used an in vitro ultrafiltration procedure [35] to prepare a fraction of u-Pl rich in molecules with nominal MW ≥50 kDa, namely the u-HMW^50kDa fraction. This was demonstrated to induce apoptosis in u-PBL, as well as in c-PBL, THP-1, and K562 mononuclear cells (Fig. 2, 3).

Proteomics data (Fig. 1) confirmed that this fraction includes large [2], or “non-dialyzable” [29], solutes, i.e., molecules exceeding the MW cutoff of dialyzer membranes used in different extracorporeal dialysis techniques and designed to prevent protein leakage (usually <30 kDa) [34, 44]. According with this, the proteomics pattern of u-HMW^50kDa showed close correspondence with that of the ultrafiltrates obtained in vivo in patients treated with PLD (nominal cutoff ≥70 kDa) [26, 38] that were originally identified to remove solutes with pro-apoptotic activity in U937 mononuclear leukemia cells [17]. Also, using 2,4-DNPH derivatization, a procedure that identifies a heterogeneous group of posttranslational modifications identified as protein carbonyls [27‒29], we confirm that u-HMW^50kDa contains the main protein decoration targets of u-Pl, with albumin (66.5 kDa) that represents by far the most abundant form [27], followed by transferrin (MW of approximately 80 kDa), fibrinogen (340 kDa), and then by a series of minor species (approx. 10% of the 2,4-DNPH-reactive material; Fig. 1).

Important enough was the finding that the main proteins and carbonylation targets of u-HMW^50kDa were also retrieved in the protein extract of u-PBL in association with increased levels of expression of the ll nonspecific multi-ligand pattern recognition receptor RAGE (Fig. 6a, b, respectively). This is a single-pass type I membrane protein and a member of the immunoglobulin superfamily expressed in different types of cells, including endothelial cells, neuronal and glial cells, neutrophils, and monocytes/macrophages. This receptor protein is involved in the cellular response to different ligands including damage-associated molecular pattern molecules (DAMPs), AGEs, and other posttranslational modifications of plasma proteins, Aβ peptides, and many others; its engagement has been demonstrated to sustain leukocyte activation, increased ROS generation, and low-grade inflammation in chronic diseases that present defects of proteostasis processes, including diabetes and uremia (reviewed in [32, 33]). A ligand-scavenging function was also described for this receptor in association with the capacity to induce programmed cell death; this was reported in macrophages in which DAMPs trigger RAGE-mediated dynamin-dependent endocytosis and pyroptosis [45] as well as in endothelial and neuronal cells in which AGEs, DAMPs, and protein aggregates can stimulate endocytosis and apoptosis (reviewed in [32]).

Therefore, the upregulation of RAGE expression and the presence of u-HMW proteins in the cellular proteome of u-PBL suggest the existence of a regulated mechanism of clearance for these large solutes by PBL endocytosis (Fig. 7). Regardless of the endocytosis mechanism (recently reviewed in [46]), the fact that uremic material from the external milieu can mobilize to mononuclear leukocytes implies the activation of retrograde transport mechanisms by the vesicular system of these cells. This is a para-physiological process that can contribute to systemic proteostasis [47, 48] involving molecular chaperones [49] active in capturing unfolded or misfolded proteins in the extracellular compartments and leading them to intracellular proteostasis modules, essentially the ER for protein repair or lysosomes and autophagosomes for proteolysis and removal/recycling [50]. Uremic toxins may hardly interfere with these processes, leading to an increased demand of proteostasis in different tissues and systems, including immune cells.

According to such a mechanistic interpretation, u-PBL showed increased levels of the UPR/ER stress response proteins PERK and IRE1-α (Fig. 6). On one hand, UPR/ERSR demonstrates the engagement of the vesicular system in these cells, and on the other hand, it can explain the increased levels of u-PBL apoptosis; in fact, the IRE1-α mediated activation of the JNK/c-Jun pathway (Fig. 6) represents a major signaling route for caspase activation and apoptotic death execution in different cell types [51], including mononuclear leukocytes [52]. u-HMW^50kDa activated the JNK pathway of these cells as well as of THP-1 monocytes (online suppl. Fig. S3). Autophagy may further contribute to the JNK-mediated apoptotic signaling of these cells (Fig. 4); in fact, autophagy is a constitutive cellular proteostasis pathway with a key role in the adaptive response to different types of cellular stresses [53], and its crosstalk with ERSR could sustain the pro-apoptotic activity of u-HMW by activating the so-called autophagy-induced apoptosis [54], a specific JNK-dependent death program active in preventing extensive cell damage and carcinogenesis, and to restore tissue homeostasis [55‒57].

Furthermore, it is conceivable to assume that the JNK activation response to u-HMW^50kDa may also link endocytosis and its effects on vesicular trafficking with the observed stimulation of ROS production (Fig. 5). Oxygen activation processes are an intrinsic component of UPR/ERSR and autophagy pathway; both the apoptotic signaling of IRE1-α/JNK pathway [58‒60] and the canonical and noncanonical pathways of autophagy [57] are redox-sensitive processes activated by an increased flux of cellular ROS [49].

An increased generation of cellular ROS represents an important apoptotic trigger in mononuclear leukocytes. We found that these reactive species mainly consist of H₂O₂, a thiol-reacting molecule acting as redox messenger of the stress response as well as of cellular proteostasis pathways [61]. In accordance with these roles of H₂O₂, we previously demonstrated that uremic solutes induce mononuclear cell apoptosis by depleting cellular thiols, a modification of the cellular redox that was corrected by the H₂O₂-scavenging agent with anti-apoptotic function N-acetyl cysteine [4].

Again, in THP-1 cells, the u-HMW^50kDa-induced generation of H₂O₂ was further enhanced after differentiation of these cells to macrophages, a cellular process that is under the influence of RAGE expression and signaling [32]. Worth of note is that the H₂O₂ produced after u-HMW^50kDa treatment in this cell line and in PBL, readily diffuses throughout the plasmalemma to reach the extracellular milieu (Fig. 5). This creates the conditions for a feed-forward mechanism of oxidative damage and leukocyte endocytosis of plasma proteins, with consequent looping into further H₂O₂ and u-HMW generation (Fig. 7).

Large uremic solutes present in the u-HMW^50kDa fraction induce abnormal endocytosis and terminal alteration of proteostasis mechanisms of mononuclear leukocytes. At the molecular level, these defects are exemplified by the cellular accumulation of plasma proteins and activation of retrograde vesicular transport mechanisms that cause JNK-mediated apoptotic signaling by terminal activation of ERSR and autophagy (Fig. 7). These alterations identify the (suicidal) role that immune cells play in an attempt to compensate for the defect of systemic proteostasis imposed by the uremic condition, a process that we here define as the “immuno-proteostasis response” (IPR) of uremia.

Since IPR might increase the risk of uremic leukopenia and immune dysfunction, its treatment is worth investigating. Possible solutions may include HD techniques and sorbents with enhanced efficacy in removing or preventing the formation of pro-apoptotic solutes included in the u-HMW^50kDa. Pharmacological therapies aimed to reduce u-HMW formation and/or IPR activation also deserve further studies, including those based on therapeutic proteostasis modulators [62, 63] that have been proposed for the treatment of the ER stress and defective proteostasis of the kidney during the development of renal disease [64].

The present study was performed in accordance with the Declaration of Helsinki and was approved by the Institutional Review Board of Fondazione IRCCS Policlinico “San Matteo” of Pavia, Italy, in 2011. This clinical trial is registered at www.clinicaltrials.gov (NCT02981992). All patients provided written informed consent. This study protocol was reviewed and approved by the IRCCS Policlinico “San Matteo” Ethical Committee located in Pavia, Viale Camillo Golgi 19, 27100 Pavia (Italy), approval number 2014-00324671.

The authors have no conflicts of interest to declare.

This work was supported by the grant program of the “Fondazione Cassa di Risparmio di Perugia” (Grant # 20420-2021.0339).

F.G., D.B., M.P., T.R., and C.L. – conception and design of the study (including clinical trial and laboratory work). D.B., M.A.G., M.P., E.C., G.G., F.G., and C.L. – acquisition, analysis, and interpretation of data. F.G., B.D., C.L., T.R., M.R., and C.R. – drafting the manuscript, revising it critically, and final approval of the version to be published.

A preprint of this article is available at Research Square (DOI: https://doi.org/10.21203/rs.3.rs-2429421/v1).