ABSTRACT
Porphyrias are caused by genetic defects in the heme biosynthetic pathway and are associated with accumulation of high levels of porphyrins that become cytotoxic. Porphyrins, due to their amphipathic nature, spontaneously associate into different nanostructures but very little is known about the effect of porphyrin speciation on the cytotoxic effects of porphyrins. Previously we demonstrated the unique ability of fluorescent biological porphyrins, including protoporphyrin IX (PP-IX), to cause organelle selective protein aggregation, which we posit to be a major mechanism by which porphyrins exerts their cytotoxic effect. Herein, we tested the hypothesis that PP-IX-mediated protein aggregation is modulated by different PP-IX nanostructures via a mechanism that depends on their oxidizing potential and protein binding ability. We demonstrate that PP-IX nanostructure formation is reversible in nature, and that nanostructure size modulates consequent protein oxidation and aggregation potential. We also show that albumin, the most abundant serum protein, preferentially binds PP-IX dimers and enhances their oxidizing ability. Additionally, extracellular albumin protects from intracellular porphyrinogenic stress and protein aggregation by acting as a PP-IX sponge. This work highlights the importance of PP-IX speciation in the context of the porphyrias, and offers insights into potential novel therapeutic approaches.
INTRODUCTION
Protoporphyrin-IX (PP-IX) is nature’s template for several essential biomolecules including heme, chlorophyll, coenzyme F430 (in methanogenic bacteria), and vitamin B12 (1). The PP-IX macrocycle consists of four pyrrole rings connected by methene bridges, and two ionizable propioniate side chains. The highly conjugated macrocycle with 18 π-electrons confers PP-IX with distinct UV-visible absorbance and fluorescence properties (2). Due to its amphiphilic nature, PP-IX tends to aggregate in aqueous solution (3). Indeed, PP-IX has been reported to exist as detergent micellarized monomers (4), dimers (5), tetramers (6), and higher order structures that are >70 kDa in size (4). The spontaneous self-association of PP-IX occurs by a combination of hydrogen-bonding (between the propionate carboxylate), and π-π stacking of the porphine ring (3,6,7). The resulting face-to-face or ‘H-oligomers have the propionate chains of adjacent PP-IX molecules in a ‘head-to-tail’ orientation (3,6,7). These supra-molecular higher order structures offer advantages such as improved responsiveness, reversibility, tunability, biomimicry, modularity, predictability, and adaptiveness, over their constituent monomers (8,9). This self-assembling property of PP-IX and similar compounds are being actively explored for a variety of applications (10–12) and synthetic porphyrin nanostructures for photodynamic therapy (13–15), photovoltaic cells (16–18), photocatalysis (19–23), and for sensor applications (23–25), have been reported. Although porphyrin macromolecular species have been extensively studied, their role in the context of protein aggregation and in the context of porphyrias is not known.
Porphyrias include eight genetic disorders that are caused by errors in any of the eight steps in the heme biosynthetic pathway (26,27). Heme biosynthesis starts in the mitochondrion where aminolevulinic acid (ALA) synthase catalyzes the rate-limiting conversion of glycine and succinyl Co-A to δ-ALA (27–30). Upon entry into the cytosol, ALA is converted in several steps, to the first cyclic tetrapyrrole, uroporphyrinogen, which is subsequently converted to copro-porphyrinogen (27–30). Coproporphyrinogen then enters mitochondria, where it is converted to heme via formation of protoporphyrinogen and PP-IX (27–30). Porphyrinogens are colorless, non-fluorescent (31) compounds, which are auto-oxidized to more stable and fluorescent porphyrins (27). Porphyrins are toxic metabolites, and their levels are tightly regulated. In porphyrias, blockages in the heme biosynthetic pathway leads to the accumulation of intermediates, with consequent organ and tissue damage (4,18,25).
Porphyrin-mediated tissue damage is proposed to occur through reactive oxygen species (ROS) generated by type I/II photosensitized reactions of porphyrins (32–34). However, the precise nature of the ROS as well as the specific targets of porphyrin-generated ROS are poorly understood (27). Recent findings demonstrated the unique properties of fluorescent porphyrins to cause organelle-selective protein aggregation though a mechanism that involves a ‘porphyrination-deporphyrination’ cycle (27,35–39). In this cycle, PP-IX binds to target proteins (i.e. porphyrination) and induces localized unfolding and conformational change (40,41). Photosensitization of protein-bound PP-IX leads to the generation of singlet oxygen (1O2), and oxidation of specific methionines to methionine sulfone or sulfoxide (27,42). Subsequent non-covalent interactions lead to protein-PP-IX lattice-like aggregate formation. During deporphyrination, acidic pH, or high salt extraction of PP-IX leads to disaggregation of the protein aggregates (27,42). We posit that this proteotoxic property of porphyrins is a major mechanism for tissue damage in porphyrias, which involves fluorescent porphyrin accumulation.
In this study, we tested the hypothesis that PP-IX-mediated protein aggregation is modulated by PP-IX speciation into supra-molecular structures, through a mechanism that involves differential oxidizing potential and protein binding.
RESULTS
pH-mediated spontaneous and reversible transformations of PP-IX nanostructures
PP-IX speciation results in distinct UV-visible and fluorescence emission signatures (3–5,43). In aqueous solution, pH and ionic strength are the principal modulators of PP-IX speciation (3). We investigated the nature of PP-IX speciation at pH 7.4 (physiological), pH 4.5 (lysosomal) and pH 9, using previously reported spectra for assignments (3–5,43). UV-visible and fluorescence spectra were collected by diluting freshly prepared PP-IX stock solutions in the indicated buffers (Fig.1A,B). In 100 mM HCl, PP-IX exists as monomers, characterized by the sharp Soret band at 409 nm (Fig.1A, Table 1). At pH 4.5, PP-IX monomers form H-aggregates, characterized by a broad Soret band with shoulders at 356 and 466 nm. At pH 9 and in 100 mM NaOH, PP-IX exists exclusively as dimers, as judged from the characteristic Soret peak at 380 nm. Notably, the absorbance spectrum at pH 7.4, shows a combination of features seen at pH 4.5, and 9, displaying a broad Soret band with slightly red shifted shoulders (379 and 469 nm) compared to the pH 4.5 spectrum (Fig1A, Table1). This suggests that at pH 7.4, PP-IX consists of a mixture of higher order aggregates and dimers. In addition to the changes in the Soret band, changes in the Q-bands in 500-700 nm region are seen. In 100 mM HCl, where the four pyrrole nitrogens are expected to be protonated, two Q-bands are observed. The number of Q bands increases to 3 (pH 4.5, 7.4), and 4 (pH 9) as the extent of protonation decreases.
The fluorescence emission spectra of the different PP-IX species (Fig.1B) revealed that a significant fluorescence quenching is associated with higher order aggregates of PP-IX. Thus, at pH 4.5, PP-IX displays a minimal fluorescence signal compared to pH 7.4 and 9 (Fig.1B, green trace).
The pH-induced PP-IX speciation is reversible (Fig.1C,D). Thus, the higher order PP-IX structures observed at pH 4.5, convert to a mixture of higher order structures and dimers (pH 7.4), and then dimers (pH 9) as the pH is increased with 5N NaOH (Fig.1C), albeit conversion is incomplete (see shoulder indicated by arrow). The transition in the reverse direction i.e. dimers → higher order structures, was observed when the pH of a PP-IX solution at pH 9 was progressively decreased to 7.4 and 4.5 with 5N HCl (Fig.1D). In earlier studies, we had reported that PP-IX induces protein aggregation in buffers containing detergents (e.g. Empigen BB, NP-40) (36,37,39,42). Given the profound effect of pH on PP-IX speciation, we tested whether detergent plays a role in porphyrin speciation. We found that irrespective of pH, 1% Empigen BB, a non-denaturing, zwitterionic detergent, shifts the equilibrium completely to monomeric PP-IX, mimicking the effect of 100 mM HCl (Fig.1E). To probe the underlying mechanism, a solution of PP-IX (10 μM) in pH7.4 phosphate buffer was supplemented with increasing concentrations of Empigen BB (Fig.1F), or NP-40, a non-ionic non-denaturing detergent, (Fig.1G) and observed the transition from higher order structures/dimers to monomers (Fig.1F-G). The absorbance at 400 nm showed a sigmoidal dependence on detergent concentration, which is characteristic of micelle formation (Fig.1H). Thus detergent-induced PP-IX monomerization appears to be caused by incorporation of monomers into detergent micelles. In summary, we observed that PP-IX reversibly transitions from dimers (at high pH) to a mixture of dimers/higher order structures to higher order structures (at low pH). Detergents and pH values of <3 favor PP-IX monomers (Fig.1I).
Smaller PP-IX nanostructures have higher oxidizing potential
Since higher order PP-IX structures show profound fluorescence quenching (Fig.1B), we characterized the effect of oligomerization on the quantum yield of PP-IX. As the nanostructure size decreases with increasing pH or with detergent, there is a significant increase in quantum yield (Φ) (Fig. 2A, supplementary Table S2). PP-IX monomers (observed in 1% Emp, pH7.4) exhibit the highest Φ.
Next, we tested whether the size-dependent increase in Φ translates to an increase in PP-IX-mediated photooxidation. For this assay, we examined PP-IX-mediated oxidative destruction using the chromophore FMN (Fig.2B). PP-IX-mediated FMN oxidation increased as a function of PP-IX concentration (Fig.2C). Importantly, the smaller PP-IX structures (monomers and dimers, observed at pH 9 and pH 7.4 + Emp), caused significantly greater FMN oxidation (Fig.2C, supplementary Table S3). In contrast a significant difference in the oxidizing capacity of PP-IX at pH 4.5 and 7.4 was not observed. These data suggest that there are two groups of PP-IX nanostructures with different oxidizing potentials. One group consists of the higher order structures that exist at pH 4.5 and 7.4, while the second group consists of monomers/dimers at pH 9 or in detergent-containing buffer (pH 7.4).
PP-IX nanostructure size modulates the type and level of PP-IX-mediated protein aggregation
Next, we tested our hypothesis that the increased oxidizing potential of PP-IX dimers/monomers leads to increased protein aggregation. PP-IX-mediated protein aggregation was clearly observed by SDS-PAGE (Fig.3A, compare lanes ± PP-IX). Specifically, the high molecular weight aggregates that failed to enter the resolving gel (Fig. 3A dotted boxes) were more prominent in the PP-IX treated samples as the pH was increased from 4.5 to 9.
Two soluble proteins, sequestome 1 (p62) and glyceraldehyde 3-phosphate dehydro-genase (GAPDH) also exhibited a dramatic increase in aggregation as the pH increases (Fig.3B,C, dotted boxes) and a corresponding loss of monomers at pH 9 (Fig.3B,C, arrowhead). PP-IX treatment caused three distinct tiers of high molecular weight aggregates in p62 (Fig.3B, dotted box). Decreasing the size of PP-IX nanostructures (via a change from pH4.5 → pH9), caused formation of heavier (with decreased electrophoretic mobility) high molecular weight aggregate. Using pH7.4+1% Empigen, we did not observe any high molecular weight aggregate for p62, although there was an evident loss of monomer (Fig.3B, lane ‘E’+PP-IX, arrowhead). We attribute the extensive loss of monomer of p62 to the loss of antibody reactivity to protein oxidation and/or aggregation following PP-IX treatment as described previously for lamin B1 (39), cyclin B1 and cdk4, (42). For GAPDH we did not observe a pronounced loss of monomer, (Fig.3D, arrowhead). In terms of high molecular weight oligomers, GAPDH showed the trend: pH9> pH7.4> 1% Emp>> pH4.5 (Fig.3D, dotted box). Notably, in contrast to p62, smaller sized PP-IX nanostructures [e.g. dimers (pH9), and monomers (1% Empigen)] lead to formation of smaller-sized high molecular weight structures (Fig3D, dotted box). In summary these results highlight the qualitative and quantitative heterogeneity in protein aggregation depending on the speciation of the PP-IX nanostructures.
Reversing PP-IX speciation reverses protein aggregation
Since the PP-IX nanostructures are interconvertible (Fig.1C,D), we hypothesized that PP-IX-mediated protein aggregation can be reversed by changing the associated PP-IX nanostructure. To test the hypothesis, we titrated PP-IX in pH 4.5 and 7.4 reaction mixtures to pH 7.4 and 4.5, respectively (Fig.3B,C, lanes 4→7 and 7→4). Notably, reversing the pH from pH 7.4 to 4.5, increased the population of unaggregated p62 and GAPDH (Fig. 3B,C), which we interpret as disaggregation due to loss of porphyrin binding. The opposite effect was observed upon changing the pH of the reaction mixture from 4.5 to 7.4.
BSA disrupts PP-IX nanostructures by preferentially binding to PP-IX dimers
We tested how PP-IX speciation modulates its binding to fatty acid-free BSA, the most abundant serum protein which has been reported to bind PP-IX (44,45). Incubation of BSA (250 μM) with PP-IX (25 μM) at pH 4.5, 7.4 and 9, increased the absorbance and fluorescence emission intensity of PP-IX, (Fig. 4A-C). In contrast, the BSA-PP-IX mixture in 1% Empigen showed no spectral changes in (Fig.4D), indicating with the absence of binding. At pH 4.5, PP-IX binding to BSA increased both the absorbance and fluorescence intensity but did not change the absorbance maxima or peak shape (Fig.4A, Table1). At pH 7.4, a dramatic shift in the Soret peak was observed in addition to increased intensity (Fig. 4B, Table1), indicating preferential binding of BSA to PP-IX dimers. The increased resolution of the Q-bands from three to four in the presence of BSA (Fig.4B, main panel and inset, Table1), suggested increased pyrrole ring nitrogen deprotonation and/or decreased mobility due to PP-IX binding to BSA. Additionally, at pH7.4 and 9 we also observed a blue shift of Q1/Q2 bands (Fig. 4C, Table1). At pH 9, an increase in intensity was accompanied by a slight red shift in the Soret peak (Fig. 4C, Table1). But Notably, the absorbance spectra of the BSA-PP-IX complex at pH 7.4 versus 9 were indistinguishable (Table 1 and Fig.S1), indicating that the same dimeric PP-IX was bound. Albumin shows perturbations of secondary structure at acidic pH, e.g. 2.4% decrease in percent helix content at pH4 versus pH7 (46). Our data suggests that despite the pH-induced structural perturbation, BSA binds PP-IX higher order nanostructures at pH4.5. In summary we conclude that: (i) there are two classes of PP-IX binding sites on BSA, one for PP-IX higher order nanostructures and a second for PP-IX dimers; ii) BSA binds preferentially to PP-IX dimers at physiological pH; and iii) BSA does not bind to PP-IX monomers.
Differential binding affinity of BSA to different PP-IX nanostructures
We used intrinsic fluorescence of BSA to monitor conformational changes induced by PP-IX binding and estimate the dissociation constant for the different PP-IX nanostructures. Since there is some pH-induced difference in the secondary structure of albumin (46), we examined the intrinsic fluorescence of BSA at pH 4.5, 7.4 and 9. The intensity of the fluorescence emission was sensitive to pH and a small red-shift was observed at pH 7.4 (Fig.5A, inset). When BSA (0.5 μM) was incubated with increasing concentration of PP-IX (0-50 μM), two types of changes were observed: a) a concentration-dependent decrease in the emission intensity (Fig. 5B-E), b) concentration dependent blue shift (Fig. 5B-F). We attribute the blue shift to the increased hydrophobicity in the environment upon PP-IX binding, with subsequent oxidative modification of aromatic residues leading to loss of fluorescence. At pH7.4 (where a mixture of dimers and higher order oligomers exist), the change in fluorescence intensity at 347 nm and the change in emission maximum showed a biphasic dependence on PP-IX concentration (Fig.5E,F). The slope of the steeper part of the curve at pH 7.4 matches that at pH 9, while the second phase matches tat at pH 4.5. This suggests that at low PP-IX concentration, dimers bind to the high affinity dimer binding site on BSA; while at higher IX concentrations, the low affinity ‘higher order structure’ binding site is occupied. To further assess this model, we analyzed the fluorescence quenching data. Only at pH 7.4 two phases were seen, consistent with two classes of binding sites (Fig. 6A-C). The number of binding sites was calculated from the slopes (Table 2), and the dissociation constant (KD) was determined by plotting Fmax-F versus PP-IX concentration (Fig. 6D). This analysis revealed two high affinity dimer binding sites and two low-affinity higher order nanostructure binding sites per BSA molecule at pH 7.4 (Table 2). Depending on the pH, the cooperativity of PP-IX binding to BSA changed.
The BSA and PP-IX complex does not dissociate after urea-induced denaturation
To test the specificity of BSA/PP-IX interaction, we examined the effect of BSA denaturation on PP-IX binding. We denatured BSA with guanidine-HCl or urea, before or after PP-IX binding. Adding PP-IX to either denatured BSA prevented PP-IX binding, as judged by the lack of absorbance or fluorescence changes (Fig. 7A,B). On the other hand, when BSA was denatured after PP-IX treatment we observed a different trend. In the presence of guanidine hydrochloride (Fig.7 C), but not in the presence of urea (Fig.7D), PP-IX was released from BSA. Guanidine-HCl purportedly affects α-helices, while urea affects β-sheets (47). Since albumin is predominantly (64%) α-helical (48), we conclude that guanidine-HCl in more efficient at disrupting its secondary structure.
PP-IX binding to BSA, enhances its oxidizing ability
When increasing concentrations of BSA (0-500 μM) were incubated with a fixed concentration of PP-IX (25 μM), we observed a concentration-dependent increase in the PP-IX Φ (Fig.8A). To test whether the oxidizing capacity of PP-IX is enhanced upon binding to BSA, the FMN oxidation assay was used. Incubating PP-IX (5 μM) with 0-5 μM BSA caused a concentration-dependent increase in PP-IX absorbance (Fig.8B). FMN oxidation was enhanced in the PP-IX+BSA mixture compared to PPI-IX alone (Fig.8C), and this was observed at all BSA concentrations (Fig. 8D). At higher BSA concentrations, there was a slight decrease in FMN oxidation, suggesting the ability of BSA to act as a sink for PP-IX generated oxidants.
PP-IX binding does not cause BSA aggregation
Based on the ability of PP-IX to induce protein aggregation [(Fig.3 and (35,36,39,42)], we tested its effect on BSA. PP-IX migrates differently in a pH dependent fashion. Higher order PP-IX nanostructures (at pH 4.5) are more prominent at the top of the gel, particularly under non-reducing conditions (Fig. 9A,D). The band corresponding to the BSA monomer became more diffuse, in the presence of PP-IX, particularly as the pH increased (Fig. 9B,E, arrowhead). This is consistent with increased BSA oxidation due to predominance of PP-IX dimers. Importantly, despite PP-IX induced BSA oxidation, there was no evidence of significant aggregation even when the gel was overloaded (Fig. 9C,F).
Unique BSA oxidation signatures based on different PP-IX nanostructures
We utilized LC-MS/MS to identify the site and type of PP-IX-mediated BSA oxidation. Since BSA shows some background oxidation, we filtered our data set and focused on oxidized residues with increased relative abundance following PP-IX treatment in three independent experimental repeats. Over 80% of oxidized residues identified were present only in PP-IX treated samples (Fig.10, boxes with black circles). As expected from the loss of BSA fluorescence emission (Fig. 5), PP-IX-led to oxidation at several tyrosine. In addition, we observed abundant protein carbonyls (+14), but no methionine oxidation (Fig.10). By contrast, methionine oxidation are abundant in other aggregated proteins including keratins and lamins (42).
While some residues (e.g., His-91, Tyr-355) were oxidized at all three pHs, the oxidation signatures showed some pH sensitive differences (Fig.10). All but one (Met-469) of the identified oxidation sites is conserved among bovine, mouse and human albumin (Met-469 is substituted by lysine in murine albumin). We propose that an overlap in the binding sites at different pHs is reflected by the ubiquitously oxidized residues, while residues that are oxidized only at pH 4.5 or 9 report on different sites and/or their level of occupancy (Fig.11).
BSA protects against intracellular porphyrin accumulation and protein aggregation after ALA+DFO treatment
Given the ability of BSA to bind PP-IX, we hypothesized that BSA modulates porphyrin-mediated protein aggregation by acting as an extracellular PP-IX sponge. To test this hypothesis, we treated HuH-7 cells with δ-aminolevulinic acid (ALA)+deferoxamine (DFO) with or without BSA added to the culture medium. ALA+DFO resulted in intracellular accumulation of PP-IX and Copro (Fig.12A,C), as well as their extracellular release (Fig.12B,D). When the medium was supplemented with 5 mg/ml BSA, significantly less porphyrin accumulated inside cells, while more porphyrin was released (Fig. 12C). We attribute this to the ability of BSA to bind PP-IX and shift the equilibrium of PP-IX secretion.
ALA+DFO treated cells with BSA in the medium also had a lower concentration of high molecular weigth protein aggregates as seen by Coomassie blue staining (Fig. 12D, dotted box). We also tested of several specific proteins including cytosolic (K8 and K18), endoplasmic reticulum (BiP and PDI), nuclear (lamins A/C and B1), and proteins involved in the degradation and clearance pathway (p62 and ubiquitin). BiP is an ER-specific HSP70 isoform and a marker for the unfolded protein response (49), and is upregulated after PP-IX induced stress (39,42). BiP is also induced when cells are treated with ALA+DFO in absence of BSA (Fig.13). Notably, all tested proteins were protected by BSA against ALA+DFO induced loss of the monomeric form and aggregation (Fig.13). We posit that albumin by acting as a ‘PP-IX sponge’, protects against intracellular porphyrin accumulation and subsequent aggregation (Fig.14).
DISCUSSION
Targeting PP-IX speciation as a potential therapeutic strategy
Porphyrin-mediated protein aggregation plays a major role in porphyric tissue damage (27,39,42). PP-IX oligomerization causes static quenching of the fluorophore (3,50), leading to dramatically reduced quantum yield, and to protein aggregation (Fig. 2A), highlighting the importance of preventing PP-IX aggregation to prevent porphyrin-mediated cell damage. Pharmacologic agents that affect PP-IX speciation could be therapeutically valuable for treating porphyria. Drugs targeting heme speciation are already known. For example, the antimalarial quinolone compounds prevent hemozoin granule formation by binding to heme monomers, and the heme monomer-quinolone complex is incorporated at the elongating hemozoin polymer, thereby inhibiting polymer elongation (51,52). Similarly, caffeine has been reported to prevent heme aggregation by forming a heme-caffeine monomeric complex (53). On the other hand, PP-IX and other related porphyrins are currently used as anti-tumor agents in photo-dynamic therapy (PDT). In the context of PDT, porphyrin-mediated protein aggregation and proteotoxicity play a beneficial cytotoxic role. For example, verteporfin a PDT agent, causes tumor selective protein aggregation (38). Thus, modulation of PP-IX speciation in a context-dependent manner could represent an attractive strategy to attenuate or enhance its toxicity.
Selectivity and specificity of PP-IX-protein interaction and related protein aggregation is guided by PP-IX speciation
A hallmark of PP-IX induced protein aggregation is its selectivity. Some known PP-IX binding proteins, such as translocator protein (TSPO) and fatty acid binding protein 1 (FABP1), do not aggregate (39). Herein we observed that BSA does not bind PP-IX monomers (Fig. 4). Thus, we posit that PP-IX speciation might be one reason underlying the selectivity of PP-IX-protein interactions. PP-IX speciation also plays a role in the nature of protein aggregation. As we demonstrate (Fig. 3), detergent micellarized-PP-IX monomers lead to qualitatively and quantitatively different protein aggregates as compared to those formed by PP-IX dimers and higher order structures. Additionally, PP-IX mediated protein aggregation is organelle selective. Proteomic analysis of protein aggregation of ALA/DFO injected zebrafish livers showed aggregation of cytosolic, nuclear, ER and mitochondrial proteins but a total absence of lysosomal protein aggregation (37). Lysosomes accumulate porphyrins similar to the cytosol, mitochondria, ER and nucleus (54–57). However, lysosomes are acidic compartments with a luminal pH of 4.5-5 (58), which aligns well with our finding that PP-IX exists as higher order and relatively inert structures with lower oxidizing and protein aggregating ability at this pH. Furthermore,, porphyrin oligomers have lower quantum yield and impaired 1O2 generation capacity (59). Indeed, live cell imaging of 1O2 demonstrated that despite lysosomal and mitochondrial accumulation of porphyrin, 1O2 was generated only in the mitochondria (60). Thus, accumulation of anionic porphyrins such as PP-IX, in the acidic environment of lysosomes (57), is not accompanied by protein aggregation. In addition to higher order structure speciation, PP-IX monomerization also modulates its toxicity as shown for detergent monomerized PP-IX which has a higher oxidizing potential (Figs. 2,3). In liver, bile salts act as surfactant detergents, and by virtue of their ability to solubilize PP-IX, play an important role in porphyrin clearance. In fact, several mouse models of protoporphyria show the important role of bile salt-PP-IX interaction in the development of cholangiopathy (61,62).
Impact of the albumin-PP-IX interaction
Albumin constitutes ~50 % of total plasma protein content (63). It is synthesized in hepatocytes then secreted into plasma (63,64) where it maintains oncotic pressure (65). In addition, albumin performs diverse functions ranging from ligand transport to serving as an antioxidant (63–66). Our data show that PP-IX binding to BSA leads to BSA oxidation and a conformational change (Figs. 5,7). Since albumin also binds to a vast network of proteins in the plasma, the so-called ‘albuminome’ (67–69), PP-IX binding could lead to disruption of components in the ‘albuminome’. In fact, oxidized human serum albumin is reported to differ significantly as compared with its un-oxidized counterpart with respect to ligand binding and antioxidant properties (70).
Porphyrin-binding proteins as a PP-IX speciation agent
Secondary structure of albumin shows considerable Binding to albumin changes the toxicity of PP-IX’s by enhancing its oxidizing ability by shifting the equilibrium from higher order to dimeric PP-IX (Fig.8,10). On the other hand, albumin acts as a ‘PP-IX sponge’, and decreases endogenous PP-IX accumulation (11–13) and protein aggregation (Fig. 13). Our findings raise the question whether albumin modulates PP-IX induced protein aggregation intracellularly in hepatocytes or extracellularly in serum. Thus, by modulating speciation, PP-IX binding proteins such as albumin, might act as novel genetic modifiers of porphyric tissue damage.
Conclusions
The reversible nature of PP-IX speciation and their different functional properties provide a potential target for therapeutic interventions. For example, our study demonstrates that PP-IX-induced protein aggregation could be reversed simply by a pH-induced shift in PP-IX speciation. This implies that transport of PP-IX-protein aggregates to the acidic lumen of the autophagolysosome can shift the equilibrium to higher order structures and lead to disaggregation of the PP-IX bound protein aggregates. Our findings highlight the dramatic difference in property and behavior of PP-IX, depending on its degree of oligomerization. Thus, PP-IX speciation could be utilized for designing better therapeutics for porphyria and improved PDT agents.
EXPERIMENTAL PROCEDURES
Cell lines and reagents
Human hepatocellular carcinoma cell line, HuH-7 cells (originally from the Japanese Collection of Research Bioresources Cell Bank) was a kind gift from Dr. Lei Yin (University of Michigan). HuH-7 cells were grown in Dulbecco’s modified Eagle’s medium (Cellgro, Manassas, VA), supplemented with 10% fetal bovine serum (Sigma-Aldrich).
PP-IX, deferoxamine mesylate (DFO), 5-ALA hydrochloride, N,N-dimethylacetamide (DMA), NaH2PO4, Na2HPO4, Flavin mono nucleotide (FMN), and ALA were obtained from Sigma Aldrich (St. Louis, MO). Fatty acid free bovine serum albumin (BSA) was from Calbiochem (San Diego, CA), coproporphyrin-III dihydrochloride (Copro) and uroporphyrin-III dihydrochloride (Uro) were obtained from Frontier Scientific (Logan UT).
Absorbance/fluorescence measurements
PP-IX absorbance and fluorescence measurements were made using a TECAN SAFIRE II microplate reader running XFLUOR4SAFIREII (Version: V 4.62n), or the Spectramax id3 running SoftMax Pro.
Quantum yield calculation
Absorbance (from 300-700 nm) and fluorescence emission spectra of 50 μM of PP-IX solution were collected. Relative quantum yield (Φ) were calculated using (Φ = ∫ Em/Abs),where ∫Em is the area under the curve for the emission spectra, and Abs is the absorbance at the excitation wavelength. ∫Em was calculated by integrating the emission spectrum using GraphPad Prismsoftware (GraphPad Software, San Diego, CA). The following excitation and emission wavelengths were used: Excitation - 437 nm, Emission - 460-850 nm (pH 4.5); Excitation - 368 nm, Emission - 400-850 nm (pH 7.4); Excitation - 389 nm, Emission - 410-850 nm (pH 9); Excitation - 410 nm, Emission - 450-850 nm (1% Empigen in pH 7.4).
FMN Oxidation
An FMN stock solution (2.2 mM) was freshly prepared in 5 mM phosphate buffer (PB), pH 7.4 and stored away from light, on ice. The FMN stock solution was diluted to a final concentration of 22 μM in 400 mM of either NaH2PO4 (pH 4.5), PB (pH 7.4), Na2HPO4 (pH 9.0), or 1% Empigen BB in PB (pH 7.4), and incubated with different concentrations of PP-IX. DMA in all the reaction mixtures was maintained at 2.8% (v/v). The control reaction mixture contained FMN (22 μM) in 2.8% DMA in pH 4.5, 7.4, or 9 buffers. After incubating the reaction mixtures (30 min, ambient light), the fluorescence emission between 500-700 nm was recorded using a SAFIRE II plate reader, after exciting at 400 nm. The percent oxidation of FMN was estimated from the loss of FMN fluorescence at 533 nm after PP-IX treatment. Additionally, where indicated, PP-IX mediated FMN oxidation was assayed as above, but in the presence of the indicated concentrations of fatty acid-free BSA. The extent of FMN oxidation was estimated by the loss of absorbance at 370 and 450 nm.
Treatment of HuH-7 cell lysates with PP-IX and biochemical analysis
Detergent-free HuH-7 cell lysates were prepared by 5 cycles of freeze-thawing then using a dounce homogenizer in 5 mM PB (pH 7.4) containing protease inhibitors cocktail (ThermoScientific, Waltham MA). The resulting solution was centrifuged (14,000 × g, 10 min, 4°C) and the supernatant and pellet were separated. The pellet was resuspended in phosphate buffered saline (PBS) supplemented with 2% SDS. Protein content in the supernatant and pellet fractions was determined using a bicinchoninic acid (BCA) assay. The final reaction mixture contained: 1 mg/ml protein with or without 50 μM PP-IX in 400 mM of either NaH2PO4 (pH 4.5), PB (pH 7.4), Na2HPO4 (pH 9.0), or 1% Empigen BB in PB (pH 7.4). Control samples had the same concentration of the vehicle, DMA. After incubating (30 min, 37°C, ambient light), the reaction was stopped by the addition of Laemmli sample buffer. Proteins (10 μg/condition) were separated by SDS-PAGE and stained with Coomassie blue or transferred to polyvinylidene fluoride (PVDF) membranes for immunoblotting. Immunoblots were visualized using horseradish peroxidase–tagged secondary antibody and chemiluminescence (Clarity Western ECL Substrate; BioRad, Hercules, CA). The antibodies used included those directed to lamin A/C, ubiquitin (Santa Cruz Biotechnology, Inc; Dallas, TX); lamin B1, p62 (Abcam; Cambridge, MA); K8 (clone TS1) and K18 (clone DC10), protein disulfide isomerase and BiP (Cell Signaling Technology; Danvers, MA).
Calculation of number of PP-IX binding site on BSA and BSA-PP-IX dissociation constant
Fatty acid-free BSA (0.5 μM) was incubated with increasing concentrations of PP-IX (0-50 μM), at pH 4.5, 7.4, and 9 buffers, respectively, for 30 minutes. Then, the solution was excited (280 nm) and the fluorescence emission spectra were collected from 320-420 nm.
Data were analyzed using a double log plot of versus log10[PP - IX], where Fmax and F are the fluorescence of BSA (347 nm) in the absence and presence of PP-IX (71). The plot yielded a straight line, and the number of binding sites was calculated using: where, K is the binding constant, and n is the number of binding sites.
The dissociation constant (KD) and Hill co-efficient (h) were calculated by plotting Fmax-F versus [PP-IX], and fitting the data to ‘Specific binding with Hill slope’ equation in GraphPad Prism. For n, KD, and h calculations, traces from each experiment were fitted individually and then averaged.
Mass spectrometric analysis of PP-IX-mediated BSA oxidation
BSA (10 μM) was treated with 500 μM PP-IX in pH 4.5, 7.4 and 9 buffers (30 min, 22°C). After treatment, 100 μl of the reaction mixture was extracted by mixing with 850 μl of methanol and 50 μl of 0.1 N perchloric acid. The resulting suspension was pelleted (20,800×g, 4°C, 20 min). The pellet was solubilized in 8M guanidine hydrochloride, 50 mM Tris HCl, pH 8.1, 20 mM DTT and incubated at 60°C, 30 min to reduce disulfide bond, followed by incubation in the dark at room temperature with 40 mM of iodoacetamide to block the reduced cysteine residues. A fraction of the sample (10%) was desalted using ziptip C4 (EMD Millipore, Burlington, MA) and dried under vacuum. The sample was treated with 0.2 μg trypsin in 10 μl of 50 mM NH4HCO3 and incubated overnight at 37°C. The digested peptides were acidified with 1 μL of 10% formic acid followed by analysis using liquid chromatography-tandem mass spectrometry (LC-MS/MS). Samples were analyzed with a Q Exactive HF tandem mass spectrometer coupled to a Dionex Ultimate 3000 RLSCnano System (Thermo Scientific) by loading onto a fused silica trap column Acclaim PepMap 100, 75 μm x 2 cm (ThermoFisher). After washing (5 min, 5 μL/min with 0.1% TFA), the trap column was brought in-line with an analytical column (Nanoease MZ peptide BEH C18, 130Å, 1.7μm, 75 μm x 250 mm, Waters) for LC-MS/MS analysis. Peptides were eluted using a segmented linear gradient from 4 to 90% B (A: 0.2% formic acid, B: 0.08% formic acid, 80% acetonitrile): 4–15% B in 5min, 15-50% B in 50 min, and 50-90% B in 15 min. Mass spectrometry data was acquired using data-dependent acquisition with a cyclic series of a full scan from 250-2000 with resolution of 120,000 followed by MS/MS (HCD, relative collision energy 27%) of the 20 most intense ions (charge +1 to +6) and a dynamic exclusion duration of 20 sec. For database search, the peak list of the LC-MS/MS was generated by Thermo Proteome Discoverer (v. 2.1) into MASCOT Generic Format (MGF) and searched against the Uniprot bovine database plus a database composed of common laboratory contaminants using an in-house version of X!Tandem (72). The search parameters were: fragment mass error (20 ppm), parent mass error (+/-7ppm), fixed modification:C-carbamidolethyl); flexible modifications: Met oxidation for the primary search and other modifications were done at the refinement stage and less than 3 modifications are set for each search. Protease specificity: trypsin (miss 2 cuts), and only spectra with loge <−2 were included in the final report. A detailed list of the oxidative modifications is listed in Supplementary Table S1.
Treatment of HuH-7 cells with ALA/DFO ± BSA
HuH-7 cells were seeded in 10 cm dishes in complete (serum containing) media. When the cells reached confluency, the media was replaced with serum-free media with ALA/DFO (1 mM/100 μM) ± fatty acid-free BSA (5 mg/ml). Control dishes had serum-free media ± fatty acid-free BSA (5 mg/ml). After 16 h, the entire volume of medium was collected and then pelleted (3000 ×g, 10 min) to remove loating cells and debris. The supernatant was saved for analysis. For the remaining adherent cells, the culture plates were rinsed twice with PBS, and the cells were harvested by scraping in 10 ml PBS. The resulting cell suspension was centrifuged (400 × g, 5 min) followed by lysis of the cell pellet with PBS supplemented with 2% SDS then analysis by SDS-PAGE. The cell lysate and culture medium were also analyzed for porphyrin content as described below.
Ultra Performance Liquid Chromatography (UPLC) analysis of porphyrins
A Waters ACQUITY UPLC system equipped with Empower software, ACQUITY H-Class PLUS (CH-A) Core, comprising of Quaternary Solvent Manager (QSM), a Sample Manager with Flow-Through Needle (SM-FTN), ACQUITY UPLC PDA Detector, ACQUITY UPLC Fluorescence Detector. A reverse-phase octadecylsilica (C18) ACQUITY UPLC BEH Shield RP18 Column, 130Å, 1.7μm, 3mm X 100mm UPLC column and ACQUITY UPLC BEH Shield RP18 VanGuard Pre-column, 130Å, 1.7μm, 2.1mm X 5mm was used. The fluorescence detector was set for excitation at 400 nm and recording emission at 635 nm. The column was eluted at a flow rate of 0.5 ml/min with linear gradients of solvents A and B (A, 0.1% formic acid in water; B, 0.1% formic acid in methanol). The solvent gradient was as follows: 0 to 1 min, 50-50% B; 1 to 2 min, 50-85% B; 2 to 3 min, 85-90% B; 3 to 9 min, 90-95% B; 9 to 9.5 min, 95-50% B; 9.5 to 14.5min, 50-50%. The cells were lysed after ALA+DFO treatment in PBS supplemented with 2% SDS. To extract porphyrins, DMA and 0.1% formic acid in methanol were added to the cell lysate or the isolated culture medium in a 1:1:2 ratio. After mixing then centrifuging (20,800 × g, 5 min) to precipitate proteins, 50 μl of the supernatant was injected into the UPLC system. Retention times of PP-IX, Uro and Copro were determined using commercially obtained standards.
Porphyrin solution preparation
PP-IX was dissolved in N,N-dimethylacetamide (1 mg/ml), while Uro and Copro were dissolved in 100 mM NaOH, to prepare 5 mM stock solutions. Care was taken not to expose the stock solution to light. The stock solutions were diluted in appropriate buffers immediately prior to use. ALA and DFO were dissolved in water to yield 500 mM and 50 mM stock solutions, respectively, then filter-sterilized, aliquoted and stored at −20°C.
Statistical Analysis
Statistical analysis of the quantum yield for different PP-IX nanostructures, FMN oxidation and porphyrin metabolite accumulation were performed using GraphPad Prism software (GraphPad Software; San Diego, CA). Statistical comparisons were carried out using ordinary one-way ANOVA analysis and Tukey’s multiple comparison test, or the unpaired T-test (2-tailed).
Author Contributions
Conceptualization (DM), Formal analysis (DM, MBO), Funding acquisition (MBO, RB, BMP), Investigation (DM, BMP, AS, HZ), Methodology (DM, HZ), Project Administration (MBO), Resources (RB), Supervision (MBO), Visualization (DM), Writing – original draft (DM), Writing – review & editing (DM, RB, MBO).
Author Disclosure Statement
No competing financial interest exists.
Supplementary Figures and Tables
ACKNOWLEDGEMENTS
This work was supported by National Institutes of Health (NIH) R01 grants DK116548 (MBO) and GM130183 (RB); Honors Summer Fellowship (College of Literature, Science, and the Arts), and Summer Undergraduate Research Fellowship (Department of Molecular and Integrative Physiology) at the University of Michigan (BMP).
ABBREVIATION LIST
The abbreviations used are:
- BSA
- bovine serum albumin
- Copro
- coproporphyrin
- DMA
- N,N-dimethylacetamide
- 1O2
- singlet oxygen
- PP-IX
- Protoporphyrin-IX
- Φ
- quantum yield
- UPLC
- Ultra Performance Liquid Chromatography
- Uro
- Uroporphyrin
- UV-Vis
- Ultra Violet-Visual spectrophotometry