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Antibodies with conformational specificity are important for detecting and interfering with polypeptide aggregation linked to several human disorders. We are developing a motif-grafting approach for designing lead antibody candidates specific for amyloid-forming polypeptides such as the Alzheimer peptide (Aβ). This approach involves grafting amyloidogenic peptide segments into the complementarity-determining regions (CDRs) of single-domain (VH) antibodies. Here we have investigated the impact of polar mutations inserted at the edges of a large hydrophobic Aβ42 peptide segment (Aβ residues 17–42) in CDR3 on the solubility and conformational specificity of the corresponding VH domains. We find that VH expression and solubility are strongly enhanced by introducing multiple negatively charged or asparagine residues at the edges of CDR3, whereas other polar mutations are less effective (glutamine and serine) or ineffective (threonine, lysine, and arginine). Moreover, Aβ VH domains with negatively charged CDR3 mutations show significant preference for recognizing Aβ fibrils relative to Aβ monomers, whereas the same VH domains with other polar CDR3 mutations recognize both Aβ conformers. We observe similar behavior for a VH domain grafted with a large hydrophobic peptide from islet amyloid polypeptide (residues 8–37) that contains negatively charged mutations at the edges of CDR3. These findings highlight the sensitivity of antibody binding and solubility to residues at the edges of CDRs, and provide guidelines for designing other grafted antibody fragments with hydrophobic binding loops.
The misfolding and assembly of peptides and proteins into prefibrillar oligomers and amyloid fibrils is linked to several neurodegenerative diseases (1,–4). To understand the contributions of polypeptide aggregation to such disorders, it is important to characterize the biochemical properties of aggregates. However, pre-amyloid and amyloid aggregates are difficult to characterize using many traditional biochemical methods that are routinely used for soluble proteins. High-resolution structural analysis of such aggregates is particularly challenging and must be performed using specialized methods such as solid state NMR (5,–13) or x-ray crystallography of small peptide fragments (14,–16). These powerful structural methods generally lack the time resolution to probe pre-amyloid intermediates and oligomers unless they can be kinetically trapped.
Given these challenges, antibodies with conformational specificity for prefibrillar oligomers and amyloid fibrils have proven valuable for biochemical characterization (17,–28). An obvious strength of such antibodies is their ability to detect specific types of aggregates formed both in vivo and in vitro. The ability of conformational antibodies to bridge in vivo and in vitro studies is important for understanding the biochemical mechanisms that contribute to protein misfolding disorders.
Several approaches have been used to generate conformational antibodies. The most widely used one is immunization, which has yielded a wide range of important antibodies specific for different types of prefibrillar oligomers and amyloid fibrils (17,–20, 25,–27). Another powerful approach that has proven useful is to employ in vitro display methods such as phage and yeast surface display (21,–24, 29,–37). These display methods afford more control over antigen presentation and have been used primarily to identify antibody fragments (rather than full-length antibodies) specific for oligomers and fibrils of several amyloid-forming polypeptides.
Nevertheless, it remains a challenge to generate antibodies with sequence and conformational specificity for different types of amyloid aggregates, which is required for many applications. To address this challenge, we are investigating the potential of designing conformational antibody fragments by mimicking the natural process of amyloid formation (38). Our approach is to graft amyloidogenic peptide segments from polypeptides such as the Alzheimer Aβ2 peptide into the complementarity-determining regions (CDRs) of single-domain (VH) antibodies. We find that these Grafted AMyloid-Motif AntiBODIES (gammabodies) recognize their cognate amyloid aggregates via homotypic interactions with modest (submicromolar to micromolar) affinity, and weakly recognize disaggregated conformers as well as amyloid aggregates formed by other polypeptides (38,–40).
The Alzheimer Aβ peptide contains two key amyloidogenic peptide segments that mediate aggregation and are structured within the amyloid core (6, 7, 41, 42). In the more amyloidogenic variant of Aβ (Aβ42), these segments typically include residues 17LVFFA21 and 31IIGLMVGGVVIA42. We posited that a VH antibody grafted with an Aβ peptide fragment containing both segments (Aβ residues 17–42) would have improved binding properties relative to shorter 10-mer variants that we have previously reported (38,–40). Moreover, our previous results (39, 40) suggested that adding hydrophilic residues to the edges of CDR3 would enhance the solubility of VH domains displaying this large amyloidogenic peptide. We also posited that these flanking polar residues would influence the conformational specificity of the Aβ VH domains as well. Therefore, we have investigated the solubility and conformational specificity of Aβ17–42 VH domains with different hydrophilic residues inserted at the edges of CDR3. We have also evaluated the generality of these findings by applying them to design antibody domains specific for islet amyloid polypeptide (IAPP), which is the polypeptide responsible for amyloid formation in type 2 diabetes.
Genes for VH expression were created using polymerase chain reaction-based gene synthesis (43) and ligated into a pET-17b bacterial expression vector (Novagen) between the NdeI and XhoI restriction sites. Restriction sites for BamHI and NotI were inserted at the edges of the CDR3 loop for introducing different sequences via ligation of synthetic primers. A PelB leader sequence was added to the N terminus of the VH gene to direct it to the periplasm. A triple FLAG tag was added to the C terminus of the VH domain, followed by a His7 tag.
Vectors containing the VH gene were transformed into BL21(DE3) pLysS cells (200132, Agilent Technologies). Transformed cells were incubated on LB-ampicillin plates for 2 days at room temperature. Next, VH expression was performed for 48 h at 20 °C and 225 rpm in 1-liter shaker flasks containing 200 ml of autoinduction media (44) supplemented with ampicillin (100 μg/ml) and chloramphenicol (35 μg/ml). Cells were sedimented by centrifugation and then discarded. Nickel-agarose resin (30230, Qiagen) was added to the clarified supernatant and incubated overnight.
The protein of interest was isolated by batch purification. The resin was washed with 100 mm imidazole at pH 7.4 to remove impurities, prior to elution with 250 mm imidazole at pH 7.4. Eluted samples were then refolded by denaturation in 6 m GuHCl at pH 7.4. Following an overnight incubation, the samples were then refolded at 4 °C by buffer exchanging into PBS, filtered, and stored at −80 °C. Refolded samples were analyzed via SDS-PAGE (WG1203BX10, Life Technologies) for samples that were reduced and boiled. The gels were then stained using Coomassie dye (24615, Thermo Scientific). The concentration of the VH domains was measured using a Bradford assay (1856209, Thermo Fisher Scientific).
VH domains were also expressed in P. pastoris. The VH genes were inserted into the pJ912 Pichia expression vector (DNA2.0, Inc.), which was linearized at the SacI restriction site to direct integration of the plasmid into chromosomal DNA at the AOX1 promoter. Cells were transformed by electroporation and then grown on plates (yeast extract peptone dextrose medium with sorbitol) supplemented with 1 mg/ml of zeocin. Colonies were picked and grown overnight in a 5-ml seed culture at 30 °C and 225 rpm. The seed cultures were then added to baffled shake flasks with 50 ml of minimal media containing glucose. The cells were grown for 60 h at 30 °C and 225 rpm, and then induced with methanol (0.5% final concentration). Expression of VH domains was detected in supernatant samples via SDS-PAGE (WG1203BX10, Life Technologies) and silver staining (LC6100, Life Technologies).
Yeast surface display was performed with the EBY100 Saccharomyces cerevisiae strain using methods that we have described previously (45). Aga2-VH fusions were subcloned from existing pET-17b constructs using PCR amplification to introduce overhangs complementary to the pCTCON2 yeast display plasmid. The PCR products were mixed with pCTCON2 plasmid that was digested with NheI and SalI, and transformed into the EBY100 strain for homologous recombination and selection on SD-CAA agar plates.
Overnight yeast cell cultures were miniprepped (D2004, Zymoprep II yeast miniprep kit, Zymo Research) and subsequently transformed into electroporation competent XL1-blue bacterial cells (200228, Agilent Technologies). Individual bacterial colonies were miniprepped (27106, Qiagen) and sequenced. Cells were then transformed with pCTCON2 plasmids encoding Aga2-VH fusions and grown overnight at 30 °C with agitation in low pH SD-CAA medium (20 g/liter of dextrose, 6.7 g/liter of yeast nitrogen base, 5 g/liter of casamino acids, 14.7 g/liter of sodium citrate, and 4.3 g/liter of citric acid) to an A600 of 1–2. Surface display of the Aga2-VH fusions was induced by pelleting the cells and resuspending them in SG-CAA media (20 g/liter of galactose, 6.7 g/liter of yeast nitrogen base, 5 g/liter of casamino acids, 8.56 g/liter of NaH2PO4·H2O, and 5.4 g/liter of Na2HPO4·2H2O). The cells were then incubated overnight at 30 °C with agitation. Both the SD-CAA and SG-CAA media were supplemented with 100 μg/ml of ampicillin and kanamycin as well as a 1× solution of penicillin/streptomycin.
Display levels were analyzed using flow cytometry. For each Aga2-VH fusion, ~106 cells were collected and washed twice with 1 ml of PBS-BX (PBS with 10 mg/ml of BSA and 1% (v/v) Triton X-100). A C-terminal epitope (myc) tag was then probed with anti-myc IgY antibody (1:200 dilution; A-21280, Life Technologies) for 1–5 h at 25 °C. After this incubation, the cells were washed with 1 ml of PBS-BX before labeling with a secondary antibody (1:100 dilution, Alexa Fluor 488-conjugated goat anti-chicken IgG, A-11039, Life Technologies) for 5 min. Fluorescently labeled cells were washed again in 1 ml of PBS-BX and the level of Aga2-VH display (as judged using the anti-myc antibody) was analyzed on a BD LSRII flow cytometer using 100,000 events.
The secondary structure of VH domains was characterized by collecting their circular dichroism (CD) spectra on a Jasco J-815 spectrometer in the far-UV region of 200–260 nm. Samples were diluted to 0.2 g/liter in PBS, and their spectra were measured using a cuvette with a 1-mm path length. Scans were collected at a rate of 50 nm/min (1 nm bandwidth and 4 s response time). For each VH domain, 10 spectra were collected and averaged. Mean residue ellipticity values were calculated from the background-subtracted spectra.
For each VH domain, the reversibility of unfolding after thermal denaturation was monitored by CD. VH domains were diluted to 0.1 g/liter in PBS in a cuvette with a 1-cm path length. Samples were heated from 25 to 95 °C at a rate of 1 °C/min, while monitoring the ellipticity at 235 nm. Following the first melt, samples were cooled to 25 °C for at least 20 min before repeating the melt. The fraction of VH domain that remained folded over the course of the melt was calculated as (θT − θU)/(θF − θU). θT is the ellipticity signal as a function of temperature, and θU and θF are the ellipticity signals for the unfolded and folded samples, respectively. The apparent melting temperature (Tm) was calculated from the initial melt.
The binding of VH domains to Protein A was measured in well plates using immobilized Protein A. Recombinant Protein A (77673, Pierce) was diluted to 5 μm in PBS, and 100 μl was added to a Nunc-Immuno 96-well plate with MaxiSorp surface coating (442404, Thermo Fisher Scientific). Protein A was incubated in the plate at room temperature overnight. Milk was then added to the plate at a concentration of 10% (w/v) and incubated at room temperature for several hours. The plate was then washed with PBS, and VH domains at varying concentrations in PBST (PBS with 0.1% (v/v) Tween 20) were added (100 μl).
The VH domains were incubated in the wells overnight at room temperature. The plate was then washed with PBS, and 100 μl of monoclonal anti-FLAG tag antibody produced in mouse (F1804-1 mg, Sigma) was added that had been diluted 1000-fold in PBST. Following a 1 h incubation of the anti-FLAG antibody, the plate was washed again with PBS, and 100 μl of an HRP-conjugated goat anti-mouse secondary antibody (32430, Pierce) was added that had been diluted 1000-fold in PBST. After 1 h of incubation, the plate was washed a final time with PBS, and 100 μl of substrate (1-Step Ultra TMB-ELISA Substrate Solution, 34028, Life Technologies) was added. The plates were developed for 1 h, and the reaction quenched by adding 2 m H2SO4 (100 μl). Signals were determined by measuring the absorbance at 450 nm for each well using a Tecan Safire2 plate reader. IC50 values were calculated from the dose-response curve that was generated for each VH domain.
The relative solubility of VH domains as a function of temperature was analyzed. Samples were diluted to ~13 μm in PBS, and divided into multiple aliquots. Each aliquot was held at a given temperature (50–95 °C) for 20 min, and then cooled to room temperature. The concentration of protein remaining in solution was measured by first centrifuging the cooled sample for 15 min at 21,000 RCF. Afterward, the top 50% of the supernatant volume was removed to determine the soluble VH concentration using the Bradford assay.
The relative solubility of VH domains at 25 °C was also analyzed by monitoring their monomeric content over a week. Samples were prepared by centrifuging them at 208,000 RCF for 1 h at 4 °C, and then recovering the top 80% of the supernatant. This procedure aimed to remove aggregated species at the beginning of the experiment. The samples were then concentrated to ~1 g/liter (~50 μm) using ultracentrifugal concentrators (10-kDa molecular weight cut-off membrane, UFC501096, Millipore). The monomeric content of the initial and 1 week samples was analyzed by size exclusion chromatography. A Waters 600F HPLC instrument was used to inject 10 μl of sample at ~1 g/liter onto a TSKgel G3000SWXL column (08541, Tosoh Bioscience) with an in-line guard column (08543, Tosoh Bioscience). The mobile phase was PBS with 0.2 m arginine (pH 7.4) at a flow rate of 0.5 ml/min. VH elution was monitored via absorbance measurements at 280 nm.
Thioflavin T, a fluorescent dye that recognizes amyloid and some non-amyloid aggregates, was added to VH samples (0.2 g/liter) to achieve a final thioflavin T concentration of 88 μm. Samples were excited at 450 nm, and emission was detected at 482 nm. The background buffer was also measured and its signal was subtracted from that of the samples. Samples were measured initially and after they had been heated to 95 °C and cooled to room temperature.
The binding of VH domains was tested for multiple antigens. These antigens include Aβ42 fibril, freshly disaggregated Aβ42 (also referred to as Aβ monomer*), IAPP fibril, and freshly disaggregated IAPP (also referred to as IAPP monomer*). Disaggregated Aβ42 was generated by first incubating the lyophilized peptide (62-0-80, American Peptide) in hexafluoroisopropanol. Following overnight incubation, the hexafluoroisopropanol was removed by evaporation. The Aβ42 peptide was dissolved in 50 mm NaOH at a peptide concentration of 1 g/liter (220 μm), and then centrifuged at 208,000 RCF for 1 h at 4 °C. The top 80% of the supernatant was recovered and passed through a 0.22-μm filter (SLGV004SL, Millipore). The sample was then diluted into acidified PBS to achieve a final pH of 7.4 and a final peptide concentration of 25 μm. Aβ42 fibrils were formed by incubating disaggregated peptide at 37 °C without agitation for at least 2 days. Prior to using the Aβ42 fibrils for binding assays, they were sonicated on ice for 3 cycles of 10 s at 100% power with 30 s of rest between cycles (FB-120 Sonic Dismembrator, Thermo Fisher Scientific).
Disaggregated IAPP was prepared by dissolving the peptide in hexafluoroisopropanol. After lyophilization, the dried peptide was dissolved in 20 mm Tris (pH 7.4) to achieve a final concentration of 32 μm. IAPP fibrils were formed by incubating disaggregated IAPP at 37 °C (300 rpm) for at least 5 days. The IAPP fibrils were sonicated on ice for 10 s at 75% power (3 times) with 30 s of cooling between rounds of sonication.
The binding of VH domains to Aβ and IAPP was characterized by an ELISA assay conducted in 96-well plates. The antigens were diluted to 2.5 μm, and 100 μl was added to a Nunc-Immuno 96-well plate with MaxiSorp surface coating (442404, Thermo Fisher Scientific) and incubated overnight at room temperature. Control wells without antigen were also included to determine the background signals. Milk dissolved in PBS (10% w/v) was then added to each well and incubated at room temperature for several hours before washing the wells with PBS. VH domains that were labeled with a biotin tag (EZ-Link Sulfo-NHS-LC-Biotin, 21335, Thermo Fisher Scientific) were diluted to varying concentrations. At each concentration, 100 μl of the VH domain sample was added to wells with immobilized antigen or background. The diluent for the VH domains was 5% (w/v) milk in PBST supplemented with 0.02% (w/v) NaN3 to reduce nonspecific interactions.
Following overnight incubation at room temperature, the biotinylated VH domains were washed from the plate. HRP-conjugated streptavidin (21126, Pierce) was diluted 30,000-fold in PBST and 100 μl was added to the wells for incubation for 1 h at room temperature. The plate was washed a final time, and 100 μl of the substrate (1-Step Ultra TMB-ELISA Substrate Solution, 34028, Life Technologies) was added to each well. The plates were developed for 1–2 h and quenched with 2 m H2SO4 (100 μl). The absorbance signal at 450 nm was measured for each well using a Tecan Safire2 plate reader. Signals were then normalized by subtracting the background value at each VH concentration, and then dividing by the background. IC50 values were calculated from the resulting dose-response curve that was generated for each VH domain.
The binding of VH domains to different antigens was also characterized using an immunoblotting assay. The antigens (Aβ42 fibril and disaggregated Aβ42, IAPP fibril, and disaggregated IAPP) were immobilized on nitrocellulose membranes (10600004, GE Healthcare) by spotting 2 μl of each antigen at different concentrations. The membranes were then blocked in 10% (w/v) milk (dissolved in PBS) for 2 h at room temperature. After the blots were washed, solutions of biotinylated VH domains (75 nm) in 5% (w/v) milk and PBST (supplemented with 0.02% w/v NaN3) were added to the blots. After overnight incubation at room temperature with mild rocking, the blots were washed three times in PBST (10 min per wash). Next, HRP-conjugated streptavidin (21126, Pierce) was added that was diluted 45,000-fold in PBST, and the blots were incubated at room temperature for 1 h. The blots were then washed again with PBST (four times for 10 min per wash) before adding the substrate (Pierce ECL Western blotting Substrate, 32106, Thermo Fisher Scientific) and imaging the blots using an x-ray film developer (Konica Minolta SRX-101A).
Site-directed mutagenesis was used to generate variants of the VH domain containing Aβ17–42 in CDR3 along with DDD at each edge of the grafted peptide. VH domains were created in which three consecutive residues in CDR3 were replaced with three alanines. The variants examined were those in which residues in the linker region of the Aβ peptide (Aβ residues 22–29) were mutated to alanine.
Site-directed mutagenesis reactions were carried out using PfuUltra II Hotstart PCR Master Mix (600850, Agilent Technologies), which was added to a mixture of the template DNA, and forward and reverse primers. The PCR involved 19 cycles of denaturation at 95 °C for 30 s, followed by 1 min of annealing at 55 °C, and 12 min of extension at 68 °C. Following the PCR, the template DNA was digested by the addition of DpnI (R0176L, New England BioLabs Inc) for 2 h at 37 °C.
To examine the effect that different types of residues have on the solubility and conformational specificity of VH domain antibodies grafted with a large amyloidogenic Aβ peptide (residues 17–42), we first cloned variants that contained the Aβ17–42 peptide flanked by three identical residues at each edge of CDR3 (Fig. 1). Variants studied included those with six identical residues that were negatively charged (DDD and EEE), positively charged (KKK and RRR), and non-charged (NNN, QQQ, SSS, TTT, and AAA).
After cloning the VH domains, we expressed them in bacteria to evaluate how different CDR3 mutations affect their final yield after expression and purification. Our previous studies revealed that increased expression of Aβ VH domains generally correlates with increased solubility (39, 40). Therefore, the relative amounts of expressed and purified VH domains can potentially be used to evaluate their relative solubility. We found that the highest expressing Aβ17–42 VH domains following purification and refolding were those with negatively charged mutations (DDD and EEE; Fig. 2).
In contrast, variants with positively charged residues (KKK and RRR) or small non-polar residues (AAA) expressed poorly (Fig. 2). SDS-PAGE analysis revealed that purified KKK and AAA VH domains could only be detected using overloaded gels, whereas the RRR variant could not be detected (data not shown). Of the VH domains with polar mutations, the TTT variant also expressed poorly (Fig. 2) and could only be detected via an overloaded SDS-PAGE gel (data not shown). Interestingly, VH domains with NNN, QQQ, or SSS mutations in CDR3 were expressed at significant levels. Although the yields of these variants were lower than the DDD and EEE variants, the ability of these non-charged residues to promote expression of the hydrophobic Aβ17–42 VH domain suggested that they increased antibody solubility.
The VH domains were next refolded in 6 m GuHCl after purification. The recovery of the VH domains after refolding was high (86–95%). Importantly, the refolding step resulted in improvements in the fraction of monomeric VH domains, especially for the NNN, QQQ, and SSS variants (data not shown).
We next sought to evaluate whether the inability to express certain Aβ17–42 VH domains was specific to our bacterial expression system or general to other expression hosts. This is a particularly important question for the KKK and RRR VH domains because their low expression in bacteria may simply be due to their inability to translocate across the cytosolic membrane (46, 47) and not due to low solubility. Therefore, we also evaluated the expression levels of the VH domains in S. cerevisiae (Fig. 3). To simplify the expression analysis, we cloned these domains as fusions to the C terminus of a yeast cell-surface protein (Aga2) and detected their display on the surface of yeast via a C-terminal myc tag.
Flow cytometry analysis revealed that the expression trends on the surface of yeast (Fig. 3) were similar to those observed for bacterial expression (Fig. 2). The expression levels of the negatively charged variants (DDD and EEE) as well as some of the non-charged variants (NNN, QQQ, and SSS) were higher than the other variants (KKK, RRR, TTT, and AAA; Fig. 3). Finally, we also evaluated expression of the KKK, AAA, and DDD variants as autonomous VH domains in P. pastoris because this host is well known to produce recombinant proteins at high levels. However, we found that expression of only the DDD variant could be detected (data not shown). These findings collectively suggest that negatively charged and some non-charged residues (asparagine, glutamine, and serine) are best at promoting expression of the hydrophobic Aβ17–42 VH domain.
We next sought to evaluate the secondary structures and stabilities of the Aβ17–42 VH domains with negatively charged (DDD and EEE) or non-charged (NNN, QQQ, and SSS) residues. We reasoned that the conformational stability of the non-charged polar variants may be higher due to the potentially unfavorable impact of the negatively charged residues on the stability of the grafted VH domains. Circular dichroism analysis revealed that the secondary structures of the charged and uncharged variants were typical of β-sheet-rich VH domains (data not shown).
To evaluate the potential impact of negatively charged mutations relative to non-charged ones on VH conformational stability, we first analyzed the relative affinity of the Aβ17–42 domains for Protein A. This analysis is motivated by the fact that folded VH3 domains (such as the Aβ17–42 domains) have a Protein A binding site on their scaffold, and the relative binding of Protein A to VH3 domains is correlated with VH stability (45, 48, 49). Using an ELISA assay to measure VH binding to immobilized Protein A, we found that the NNN, QQQ, and SSS variants had similar IC50 values of ~1–2 nm, whereas the DDD and EEE variants had higher IC50 values (~5–10 nm; Fig. 4 and Table 1). The higher IC50 values for the negatively charged variants suggested they are less stable than the other polar variants, which may be due to intramolecular repulsion.
We further examined the conformational stability of the Aβ17–42 VH domains by measuring their apparent melting temperature (Tm) values using circular dichroism. The DDD and EEE variants had lower Tm values (~69 and ~71 °C, respectively) than the NNN, QQQ, and SSS variants (~75 °C; Fig. 5 and Table 1). These findings are consistent with the Protein A results (Fig. 4) and also suggest that the VH domains with negatively charged mutations in CDR3 are less conformationally stable than the variants with non-charged mutations.
We have previously found that negatively charged mutations in CDR3 of grafted VH domains prevent irreversible unfolding of some hydrophobic VH domains but not others (39, 40). Thus, we sought to evaluate if the DDD and EEE variants promoted reversible unfolding and refolding of the Aβ17–42 VH domains as well as to what extent non-charged mutations could promote such reversible unfolding. Circular dichroism was used to monitor the ellipticity of the samples at a fixed wavelength (235 nm) as a function of temperature. The DDD and EEE variants showed largely reversible unfolding and refolding behavior (Fig. 5), consistent with our previous findings (39, 40, 50). Surprisingly, the NNN variant also displayed unfolding behavior that was largely reversible, whereas the QQQ and SSS variants unfolded irreversibly. These differences were also apparent in terms of the circular dichroism spectra at 25 °C for VH domains before and after being heating to 95 °C. The DDD, EEE, and NNN VH variants largely regained their initial secondary structure after being heated to 95 °C and cooled, whereas the secondary structure of the QQQ and SSS variants was significantly perturbed (data not shown).
These results suggested that the QQQ and SSS VH domains aggregated at high temperature. To investigate this in more detail, we performed solubility measurements after heating the Aβ17–42 VH domains to different temperatures (50–95 °C) and cooling them to 25 °C (Fig. 6). The solubility of the DDD and EEE variants was high and unaffected by heat stress (Fig. 6A). The NNN VH variant was modestly less soluble than the DDD and EEE variants, but significantly more soluble than the QQQ and SSS variants. These solubility differences appear related to the formation of amyloid-like aggregates (Fig. 6B). The QQQ and SSS VH domains displayed high thioflavin T fluorescence after heating, whereas the other polar variants displayed similar (DDD) or modestly increased (EEE and NNN) thioflavin T fluorescence after heating.
We also investigated the relative solubility of the Aβ17–42 VH domains at room temperature (25 °C) and a higher antibody concentration (~50 μm) over the course of a week (Fig. 7) to evaluate similarities or differences with the non-native solubility data (Fig. 6). Samples were evaluated for their fractional monomer content using size exclusion chromatography. The monomer content of the DDD and EEE variants remained constant at ~90% over the course of 1 week at 25 °C. The NNN variant had a lower initial monomer content (~65%), which remained roughly constant after 1 week. The other variants that initially contained 66% (QQQ) and 53% (SSS) monomer displayed further aggregation over the course of 1 week. These findings reveal that the relative native and non-native solubilities of the Aβ17–42 domains display similar trends, which is consistent with our hypothesis that the solvent-exposed Aβ17–42 peptide in CDR3 mediates aggregation of both unfolded and folded VH domains.
To evaluate the generality of our findings, we also sought to design a grafted VH domain specific for IAPP. IAPP is extremely amyloidogenic and several small peptide segments derived from residues 8–37 are known to form fibrils (51,–55). Thus, we chose to graft this large peptide segment (residues 8–37) into CDR3 along with negatively charged residues (DED) at each edge of CDR3. This IAPP8–37 VH domain expressed well (94 mg/liter) and showed similar secondary structure as the DDD Aβ17–42 VH domain (data not shown). Upon thermal denaturation, it was also able to unfold in a nearly reversible manner (Fig. 8A), its melting temperature (71 °C) was similar to those of the DDD and EEE Aβ17–42 VH domains (69–71 °C), and it was largely monomeric like the DDD Aβ17–42 VH domain (Fig. 8B). In contrast, the same IAPP8–37 VH domain with NNN residues at each edge of CDR3 showed reduced expression (15 mg/liter), suggesting that the negatively charged residues enhanced solubility of this antibody domain.
The significant effect of the polar mutations flanking the Aβ17–42-grafted peptide on VH solubility led us to also evaluate how such mutations influence antigen binding. Using an ELISA assay to measure concentration-dependent binding, we found that the DDD, NNN, QQQ, and SSS variants all recognized Aβ42 fibrils. The EEE variant was not evaluated in detail because it displays similar binding activity as the DDD variant (data not shown). The fibril IC50 values were lowest for the DDD (43 ± 15 nm) and NNN (43 ± 12 nm) variants, and highest for the QQQ (114 ± 75 nm) and SSS (346 ± 64 nm) variants (Fig. 9A and Table 2). Interestingly, the most significant difference between the Aβ17–42 VH domains was their ability to recognize disaggregated Aβ. The negatively charged (DDD) variant showed little binding to disaggregated Aβ42 (Fig. 9A). In contrast, VH variants with non-charged mutations were able to bind to disaggregated Aβ42. The SSS and QQQ variants displayed increased binding signals for disaggregated Aβ relative to fibrils. The IC50 values for binding of the NNN, QQQ, and SSS variants to disaggregated Aβ were similar (200–300 nm; Table 2).
Given that some of the VH domains form higher order species (Fig. 7A), we evaluated whether such multimers may influence the observed conformational specificity or IC50 values of the Aβ17–42 VH domains. To accomplish this, we enriched the fraction of VH monomer in the purified samples for the QQQ and SSS VH domains and evaluated their binding. A rigorous centrifugation step (208,000 RCF for 1 h) reduced the fraction of high molecular weight species from 15 ± 1 to 9 ± 1% for the QQQ VH domain and from 17 ± 1 to 11 ± 1% for the SSS VH domain.
For the QQQ and SSS VH domains, samples with reduced aggregate content had IC50 values that were within error of the control samples for binding to disaggregated Aβ (data not shown). This suggests that the presence of VH multimers did not significantly affect binding. Interestingly, we observed that the reduction of VH multimers modestly enhanced the normalized binding signals (data not shown). Collectively these findings suggest that the enhanced binding of the QQQ and SSS VH domains (relative to the DDD variant) to disaggregated Aβ is not strongly linked their higher aggregate content.
We also found that the polar residues at the edges of CDR3 influenced the sequence specificity of the Aβ17–42 VH domains. The DDD variant displayed high specificity for Aβ fibrils relative to IAPP fibrils (Fig. 9A). The NNN, QQQ, and SSS variants bound preferentially to Aβ fibrils but also displayed modest binding to IAPP fibrils. We confirmed that the specificity of the Aβ17–42 VH domains for Aβ fibrils was not simply due to low reactivity of IAPP fibrils by evaluating binding of the IAPP8–37 VH domain (Fig. 9B). This VH domain with DED residues at the edges of CDR3 recognized IAPP fibrils (IC50 value of 126 ± 68 nm) and weakly bound to either Aβ conformer.
We also evaluated the binding activity of the Aβ and IAPP VH domains using an immunoblotting assay (Fig. 10). Aβ42 and IAPP fibrils and disaggregated peptides were deposited on nitrocellulose membranes at a range of concentrations and probed with VH domains as well as with conventional antibodies. The DDD Aβ17–42 VH domain displayed significant conformational specify for Aβ fibrils relative to disaggregated Aβ and little cross-reactivity with IAPP conformers. The NNN, QQQ, and SSS variants displayed similar binding to fibrils. However, they also weakly recognized disaggregated Aβ (Fig. 10), which is generally consistent with the ELISA results (Fig. 9). Moreover, the background signals for the NNN VH domain and especially for the QQQ and SSS VH domains were higher than for the DDD variant. This suggests that the non-charged variants were stickier and less well suited for immunoblotting applications. Interestingly, the specificity of conventional antibodies (LOC and OC) with conformational specificity for Aβ fibrils was only observed at the lowest amounts of deposited Aβ (0.5 pmol). Moreover, these antibodies that recognize fibrils of different amyloid-forming polypeptides (19, 56, 57) either failed to recognize IAPP fibrils or recognized both IAPP conformers after long exposure times (LOC, data not shown). As expected, conventional antibodies that recognize sequence-specific epitopes within Aβ42 (NAB 228, 12F4, and 4G8) recognized both Aβ fibrils and disaggregated Aβ.
Conversely, we observed that the IAPP8–37 VH domain selectively recognized IAPP fibrils relative to disaggregated IAPP, and showed low cross-reactivity with Aβ conformers (Fig. 10). This is in contrast to a conventional IAPP sequence-specific antibody (R10/99) that recognized both IAPP conformers. These results collectively demonstrate that VH domains grafted with amyloidogenic peptides can be generated with sequence and conformational specificity for amyloid fibrils.
The fact that the DDD variant of the Aβ17–42 VH weakly recognizes disaggregated Aβ relative to the other Aβ VH variants without charged mutations suggests that repulsive electrostatic interactions may mediate the conformational specificity of the DDD variant. This is further supported by the fact that Aβ42 is predicted to have an acidic isoelectric point (~5) and contains six negatively charged residues, some of which are adjacent to the most amyloidogenic regions. To test this hypothesis, we compared the binding of the DDD Aβ17–42 VH to Aβ and IAPP conformers at normal (0.14 m) and elevated (1 m) concentrations of sodium chloride (Fig. 11A). Importantly, this VH domain displayed significantly enhanced binding to disaggregated Aβ at high salt, and did not show increased binding to IAPP conformers. The DDD Aβ17–42 VH also displayed modest increases in its binding signals for Aβ fibrils.
We also evaluated whether the enhanced binding of the DDD antibody variant to Aβ at high salt was mediated by salt-induced VH oligomerization. To test this possibility, we used size exclusion chromatography to analyze the potential oligomerization of the DDD Aβ17–42 VH as a function of salt concentration. However, the levels of the high molecular weight species of the DDD VH domain were a weak function of salt (5% at 0.14 m NaCl, 6% at 1 m NaCl). Thus, we conclude that the enhanced affinity of the Aβ17–42 VH for disaggregated Aβ is likely due to the screening of repulsive electrostatic interactions rather than salt-induced oligomerization of the VH domain.
The enhanced binding for the DDD Aβ17–42 VH to Aβ42 monomer (pI ~5) at high salt suggests that repulsive electrostatic interactions influence the corresponding conformational specificity. In contrast, IAPP is a basic polypeptide (theoretical pI of >10) with only three residues that potentially could be charged (Lys1, Arg11, and His18) and which are peripheral to the most amyloidogenic region (IAPP residues 22–27). Thus, we expected that the binding of the IAPP8–37 VH with negatively charged residues (DED) at the edges of CDR3 would be weakly impacted by changes in salt concentration. Indeed, we find that high salt does not alter IAPP8–37 VH binding to IAPP or Aβ conformers (Fig. 11B). These results reveal important differences in the mechanisms of conformational specificity for the Aβ and IAPP VH domains.
We next generated and analyzed a series of alanine mutations in CDR3 of the DDD Aβ17–42 VH to understand how residues within the grafted Aβ peptide modulate antibody expression and binding. We focused on Aβ residues 22EDVGSNKG29 that link the two major aggregation-prone regions (Aβ residues 17LVFFA21 and 31IIGLMVGGVVIA42) within Aβ42. Mutants were generated in which three consecutive residues were replaced by three alanines.
We find that Aβ residues 21AED23 are critical for expression, as mutating them to alanines significantly reduced VH expression (Fig. 12A). Likewise, mutation of residues 27NKG29 to three alanines greatly reduced expression. However, mutation of Aβ residues 23DVG25 or 25GSN27 to three alanines resulted in Aβ17–42 VH domains with similar or modestly lower expression levels relative to wild-type VH (Fig. 12A). Size exclusion chromatograms and far-UV CD spectra for the mutants and wild-type were also similar (data not shown). Their binding behavior, however, was significantly different (Fig. 12B). The 23AAA25 and 25AAA27 VH mutants bound to disaggregated Aβ better than wild-type. These findings indicate that hydrophilic Aβ residues 22–29 generally reduce Aβ self-association and increase solubility, and that Aβ residues 23–27 reduce Aβ17–42 VH binding to disaggregated Aβ.
Our studies aimed at designing anti-amyloid antibody fragments were originally motivated by a previous report using a similar motif-grafting approach to generate full-length antibodies specific for aggregated forms of the mammalian prion protein (PrP) (58). These antibodies were generated by grafting PrP peptides containing residues 89–112 (24-mer) or 136–158 (23-mer) into heavy chain CDR3 of an antibody originally specific for an HIV protein. These peptides were identified based on their importance in mediating conversion from PrPC to PrPSc (29, 59,–63). The resulting grafted antibodies recognized the aggregated, infectious form of PrP (PrPSc) but not the soluble, non-infectious form (PrPC).
Notably, the grafted PrP peptides each contain four positively charged residues (58, 64). Mutating these residues to alanine eliminates binding for the grafted variant with PrP residues 89–112 in heavy chain CDR3 (this analysis was not reported for the other variant). A third PrP peptide (residues 19–33) was later identified that also mediates binding to aggregated forms of PrP when grafted into heavy chain CDR3 (64). This peptide also contains four positively charged residues and binding is eliminated when such residues are mutated to alanine. Collectively these results suggest that the binding mechanism of these PrP conformational antibodies involves electrostatic interactions.
Our results suggest that Aβ and IAPP-grafted antibody domains employ non-electrostatic interactions to mediate binding to amyloid aggregates. This is supported by the fact that fibril binding for the Aβ17–42 and IAPP8–37 VH domains with negatively charged CDR3 mutations is similar or better at high salt than at low salt (Fig. 11). Moreover, fibril binding for Aβ17–42 VH domains was similar for CDR3 mutations that have side chains of similar size (aspartate and asparagine; Fig. 9).
Nevertheless, we find that repulsive electrostatic interactions are important for determining the affinity of Aβ17–42 VH binding to disaggregated Aβ. Addition of negatively charged residues (DDD) at the edges of the grafted Aβ17–42 peptide reduced VH binding to disaggregated Aβ, whereas removal of negative charge in the grafted Aβ17–42 peptide (23DVG25 to 23AAA25) increased VH binding. These findings suggest that the conformational specificity of the Aβ17–42 VH for binding to Aβ fibrils relative to disaggregated Aβ is strongly mediated by repulsive electrostatic interactions.
Collectively our results suggest that the mechanism for the conformational specificity of the DDD Aβ17–42 VH domain is due to multiple contributions. One factor may be the entropic penalty for binding of the grafted Aβ17–42 peptide to disaggregated Aβ, the latter of which is largely unstructured. We reason that the interaction of this grafted VH domain with Aβ fibrils is more favorable because of the lower entropic penalty for binding to structured fibrils. A second factor that appears to contribute to the conformational specificity of the DDD Aβ17–42 VH domain is electrostatic repulsion between negatively charged residues in CDR3 and those within Aβ. We speculate that the DDD variant binds to Aβ fibrils because amyloidogenic (homotypic) interactions involving Aβ residues 17–42 are more significant than the repulsive electrostatic interactions that discourage binding to disaggregated Aβ. It is also possible that the effective net charge of Aβ within structured fibrils is lower than for disaggregated Aβ, which may also contribute to the observed conformational specificity. Moreover, avidity effects due to the polyvalent nature of aggregates relative to monomers may also contribute to the observed conformational specificity (65).
Our findings for the DED IAPP8–37 VH domain provide further insight into the mechanisms of conformational specificity for such grafted antibodies. Notably, this VH domain binds well to IAPP fibrils and poorly to disaggregated IAPP. The fact that the IAPP peptide contains little charge and that the conformational specificity of the DED IAPP8–37 VH domain is insensitive to salt concentration suggests that its binding mechanism is largely mediated by non-electrostatic interactions. The conformational specificity of this VH domain may be due to the reduced entropic penalty for binding to structured fibrils relative to disordered peptide and/or avidity effects due to polyvalent aggregates. These speculative conclusions will need to be tested for additional grafted VH domains and using different types of aggregates.
The fact that the expression levels of the Aβ17–42 VH domains were strongly influenced by elimination of charged residues in the grafted Aβ17–42 peptide also deserves further consideration. Our alanine mutational data revealed that Aβ residues 22ED23 are critical for expression, which is consistent with previous studies that have highlighted the importance of these residues for Aβ solubility (66,–68). Deletion or substitution of these negatively charged residues for non-charged or positively charged residues, as occurs for Aβ variants associated with familial forms of Alzheimer disease, has been found in some cases to significantly increase the aggregation rates of Aβ (69,–71).
Our findings related to the impact of CDR3 mutations on VH solubility also deserve further consideration. We have previously shown that negatively charged mutations are effective at increasing the solubility of VH domains grafted with shorter (10-mer) amyloidogenic peptides (39, 40), which is consistent with our findings here. Moreover, we have also previously found that positively charged mutations are less effective at increasing expression and solubility of grafted VH domains unless the antibody scaffold has a highly basic isoelectric point (39). Our current findings for a scaffold with a near-neutral isoelectric point (theoretical pI of ~8.5 when omitting the CDRs and FLAG tags) are consistent with our previous observations given the low expression levels of the Aβ17–42 VH domains with positively charged mutations in both Escherichia coli and P. pastoris, as well as the low display levels of such variants on the surface of S. cerevisiae.
It is notable that the polar residues asparagine, glutamine, and serine enabled VH expression at sufficient levels for purification and analysis, whereas threonine did not. Moreover, asparagine was found to be most solubilizing, followed by glutamine and serine. We originally posited that serine would be the most solubilizing of these polar residues for multiple reasons. First, we suspected that the propensity of polyglutamine and polyasparagine to self-associate and aggregate (72, 73) would make these residues less solubilizing than multiple serines when introduced into CDR3. Moreover, previous work by others demonstrated that serine is the most solubilizing non-charged residue when introduced into a solvent-exposed loop within Ribonuclease Sa (74). However, there are several key differences between this previous study and our study, including the number of mutations (one for Ribonuclease Sa and six for VH domains) and the solution conditions (1–2 m ammonium sulfate at pH 4.25 for Ribonuclease Sa and PBS at pH 7.4 for our VH domains) that make it difficult to compare them.
Nevertheless, our findings are consistent with several hydropathy indices that report asparagine as the most hydrophilic non-charged residue followed by glutamine, serine, and threonine (in that order) (75, 76). Thus, it is possible that residues such as asparagine and glutamine are simply increasing the local hydrophilicity adjacent to the grafted hydrophobic peptides and thereby increasing VH solubility. Moreover, it is likely that six asparagine or glutamine residues in CDR3 are unable to mediate the aggregation observed for longer repeat lengths of polyasparagine and polyglutamine (73, 77). It is also notable that hydropathy indices do not describe the overall solubility trends we observed for charged and non-charged residues. In particular, positively charged mutations are typically predicted to be more hydrophilic than non-charged polar residues and, in some cases, more hydrophilic than negatively charged residues (75, 76).
We have also identified a number of factors that influence the performance of our grafted VH domains. First, we find that the longer grafted Aβ (26-mer) and IAPP (30-mer) peptides lead to improved binding relative to the shorter (10-mer) grafted peptides that we have previously reported (38,–40, 78). Our findings as well as those for PrP antibodies (58, 64) suggest that peptide lengths of 23–30 residues are a logical starting point for designing antibodies and antibody fragments specific for other amyloid-forming polypeptides. Second, we find that our VH domains display some propensity to interact non-specifically with well plates and immunoblotting membranes unless the experiments are performed in milk. Thus, it is important to block well plates and membranes with 10% milk, and conduct the binding experiments in at least 5% milk using our VH domains. This also suggests that the Aβ and IAPP VH domains are not appropriate for use in assay formats that are incompatible with milk.
We also find that the detection sensitivity is higher using biotin-based detection (streptavidin-HRP) than peptide tag-based detection (anti-FLAG and secondary antibody-HRP). Thus, it is recommended to biotinylate the VH domains at low levels (~4–7 biotins per domain) to maximize specificity and detect them using standard biotin-specific reagents. Moreover, we find that sequence specificity of the VH domains is reduced as the amount of immobilized antigen is increased in immunoblotting assays. For the NNN, QQQ, and SSS Aβ17–42 VH domains, we observe some cross-reactivity with IAPP fibril in well plates at VH concentrations >100 nm (Fig. 9). We also sometimes observe cross-reactivity of the Aβ17–42 (DDD) and IAPP8–37 (DED) VH domains with non-cognate antigens such as α-synuclein and Tau for immunoblots loaded with >200 ng of protein (data not shown). Thus, it is important to evaluate the binding of such VH domains over a range of amounts of immobilized antigen to identify suitable conditions for maximizing sequence specificity.
We also find that the assay format influences the extent of conformational specificity of the VH domains. The Aβ17–42 (DDD) and IAPP8–37 (DED) VH domains show significant conformational specificity both in well plates and immunoblot assays (Figs. 99–11). However, some differences were observed for the QQQ and SSS Aβ17–42 VH domains. The higher binding signals of these domains for disaggregated Aβ in the well plate assay and for fibrils in the immunoblotting assay may be due to the different mechanisms of antigen immobilization and/or the impact of varying antigen concentration (immunoblotting) versus varying antibody concentration (well plates). Our findings demonstrate the importance of standardizing assays with well defined fibril and disaggregated peptide samples to properly interpret the results. Indeed, we find that conventional antibodies with conformational specificity for fibrils (LOC and OC) only show fibril specificity at low amounts of immobilized antigen (Fig. 10). Interestingly, we also observe a similar level of conformational specificity for a sequence-specific Aβ antibody (NAB 228) at low amounts of immobilized antigen. This demonstrates that the observed conformational specificity of many antibodies (including conventional ones) is dependent on assay conditions and must be interpreted carefully (65).
A key next step will be to identify the residues within the grafted Aβ17–42 and IAPP8–37 peptides that contribute most to the affinity and conformational specificity of the VH domains. This is important to further optimize the antibody domains as well as to guide the design of anti-amyloid antibodies specific for other amyloid-forming polypeptides. We are currently expanding our use of alanine-scanning mutagenesis to the entire CDR3 to identify the most important residues for expression, solubility, and binding. It will also be important to further increase the affinity and conformational specificity of these grafted domains using directed evolution methods. We are currently randomizing positions within and near CDR3 of similar grafted VH domains, and selecting variants with improved affinity and conformational specificity. We expect that these and related approaches will lead to the rapid design and optimization of antibodies with high conformational and sequence specificity for amyloid aggregates.
C. C. L and P. M. T. designed the research; C. C. L., M. C. J., K. E. T., F. M., and S. E. D. performed the experiments; R. A. and D. P. R. provided reagents, technical advice, and comments on the manuscript; and C. C. L., M. C. J. and P. M. T. wrote the paper.
We thank members of the Tessier lab for their helpful suggestions on our manuscript.
*This work was supported, in whole or in part, by National Institutes of Health Grants R01GM104130 (to P. M. T.) and R01GM078114 (to D. P. R.), National Science Foundation CBET Grants 0954450 and 1159943 (to P. M. T.), grants from the Pew Charitable Trust (Pew Scholars Award in Biomedical Sciences (to P. M. T.), New York Capital Region Research Alliance (to P. M. T.), and Richard Baruch M.D. Chair (to P. M. T). P.M.T. has received consulting fees and/or honorariums for presentations of this and/or related research findings at MedImmune, Eli Lilly, Bristol-Myers Squibb, Janssen, Merck, Genentech, Amgen, Pfizer, Adimab, Abbvie, Roche, Boehringer Ingelheim, Bayer, Abbott, DuPont, Schrödinger, and Novo Nordisk. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.
2The abbreviations used are: