High‐efficient bacterial production of human ApoA‐I amyloidogenic variants

Rita Del Giudice; Jens O Lagerstedt

doi:10.1002/pro.3522

. 2018 Nov 15;27(12):2101–2109. doi: 10.1002/pro.3522

High‐efficient bacterial production of human ApoA‐I amyloidogenic variants

Rita Del Giudice ^1,^✉, Jens O Lagerstedt ^1,^✉

PMCID: PMC6237697 PMID: 30291643

Abstract

Apolipoprotein A‐I (ApoA‐I)‐related amyloidosis is a rare disease caused by missense mutations in the APOA1 gene. These mutations lead to protein aggregation and abnormal accumulation of ApoA‐I amyloid fibrils in heart, liver, kidneys, skin, nerves, ovaries, or testes. Consequently, the carriers are at risk of single‐ or multi‐organ failure and of need of organ transplantation. Understanding the basic molecular structure and function of ApoA‐I amyloidogenic variants, as well as their biological effects, is, therefore, of great interest. However, the intrinsic low stability of this type of proteins makes their overexpression and purification difficult. To overcome this barrier, we here describe an optimized production and purification procedure for human ApoA‐I amyloidogenic proteins that efficiently provides between 46 mg and 91 mg (depending on the protein variant) of pure protein per liter of Escherichia coli culture. Structural integrity of the amyloidogenic and native ApoA‐I proteins were verified by circular dichroism spectroscopy and intrinsic fluorescence analysis, and preserved functionality was demonstrated by use of a lipid clearance assay as well as by reconstitution of high‐density lipoprotein (HDL) particles. In conclusion, the use of the described high‐yield protein production system to obtain amyloidogenic ApoA‐I proteins, and their native counterpart, will enable molecular and cellular experimental studies aimed to explain the molecular basis for this rare disease.

Keywords: apolipoprotein A‐I, high‐density lipoprotein, HDL, amyloidosis, high‐yield protein production

ABBREVIATIONS

ApoA‐I: apolipoprotein A‐I
CD: circular dichroism
DMPC: 1,2‐dimyristoyl‐sn‐glycero‐3‐phosphocholine
FC: free cholesterol
HDL: high‐density lipoprotein
IPTG: isopropyl‐beta‐thiogalactopyranoside
PBS: phosphate buffer saline
POPC: 1‐palmitoyl‐2‐oleoyl‐sn‐glycero‐3‐phosphocholine
rHDL: reconstituted HDL
TEV: tobacco etch virus
WT: wild‐type

Introduction

Protein and peptide instability, and their propensity to aggregate into organized fibrils or amyloids, is the underlying cause for common and rare conditions collectively named amyloid diseases.1, 2 Depending on the identity of the associated amyloidogenic protein or peptide, their accumulation into amyloids may lead to neurological disorders, due to protein deposition in the brain, as in the case of Alzheimer disease or Parkinson's disease, metabolic disorders, caused by deposits in pancreas, as in Type 2 diabetes, or to systemic conditions with deposits in several different vital organs, including heart, liver and kidney. The range of the physiological effects that may be experienced by the affected individuals includes cognitive impairment, metabolic imbalance, and ultimately organ failure.1, 2

The depositions of proteins into amyloids in the affected organs are typically a slow process that occurs over decades with evident clinical manifestation only later in life. Enabling identification and treatment of this class of diseases at an early stage is, therefore, critical to prevent disease progression and severity. Detailed studies of the protein/peptide behavior at the molecular level are thus needed and will facilitate the development of novel therapeutics for these patient groups. To achieve this, biochemical, biophysical and imaging approaches including small‐angle X‐ray scattering,3 cryo electron microscopy,4, 5 atomic force microscopy,6 electron paramagnetic resonance spectroscopy,7 synchrotron radiation circular dichroism (CD),8, 9 and solid‐state nuclear magnetic resonance spectroscopy10 have been used. To perform these types of analyses, proteins or peptides must be isolated with high quality and at sufficient amounts. However, for amyloidogenic proteins that are prone to aggregate, obtaining the required protein levels and qualities commonly constitutes a major challenge. Much effort has therefore to be invested into the establishment of experimental protocols for the high‐efficient production and purification of amyloidogenic proteins.

Apolipoprotein A‐I (ApoA‐I) is the main protein component of high‐density lipoprotein (HDL) and, as such, it has a central role in the uptake and transport of cholesterol and lipids in circulation11, 12 and on glucose control.13, 14 ApoA‐I is, therefore, considered to be beneficial to our health by reducing the risk for cardiovascular diseases11, 15, 16 and by counteracting hyperglycemia.17 However, specific mutations in the APOA1 gene, and the consequent single amino acid substitutions in the translation products, are associated with protein instability and progressive accumulation of protein fibrils in the affected organs, such as heart, liver, kidneys, skin, nerves, ovaries, or testes, eventually leading to organ failure.18, 19 Interestingly, amyloidogenic mutations in the Residues 1–75 of the 243‐amino acid long ApoA‐I protein (e.g., the G26R and L75P variants) lead mostly to hepatic, renal and testis amyloidosis, whereas mutations in Residues 173–178 (e.g., the L174S and L178H variants) mainly lead to cardiac, laryngeal and cutaneous protein accumulation.20, 21, 22, 23 While several advances have been made to understand the low‐resolution structural characteristics of amyloidogenic ApoA‐I variants,7, 8, 24, 25, 26 much remains to be explained regarding the molecular mechanisms driving the ordered aggregation process, the variant‐specific tissue preferences, as well as how the aggregation can be therapeutically prevented. Here, we have evaluated different cell growth conditions, in terms of culture medium formulations and temperature during the induction of protein expression, in order to establish a new production protocol to obtain high yields of the aggregation‐prone ApoA‐I variants. This novel protocol will enable detailed biochemical and biophysical studies to unravel the molecular mechanisms in hereditary systemic ApoA‐I amyloidosis.

Results and Discussion

Amyloidogenic proteins are usually instable and prone to form aggregates, which makes heterologous expression and purification very challenging. The amyloidogenic ApoA‐I variant L75P, which is characterized by a relaxed tertiary structure and by a very low stability, is an example of this.24 Yet, the availability of highly efficient expression systems that generate sufficient amounts of protein are crucial in order to perform accurate functional and structural studies on the molecular mechanism underlying ApoA‐I amyloidosis.

Effect of medium composition and temperature on the expression levels of ApoA‐I proteins

In order to find the most suitable expression system for the high‐yield production of the L75P‐ApoA‐I variant, we evaluated different media formulations and also investigated the effects of two different temperatures during the gene expression induction phase. Media with different compositions and complexities, such as LB, TB, and NZYM in the presence or absence of 0.2% glucose as carbon source, were used in the analyses.

Escherichia coli cells, transformed with the vector encoding either WT or L75P‐ApoA‐I, were grown in the different media at 37°C. When bacterial cultures reached OD_600nm of 0.5–0.7, protein expression was induced by the addition of IPTG and cells incubated at either 37°C (Fig. 1, left panels) or 30°C (Fig. 1, right panels). As shown in Figure 1, ApoA‐I WT was efficiently produced in all the conditions tested, with higher expression levels in NZYM broth in the presence of glucose and at 30°C. Notably, L75P‐ApoA‐I expression was detected only, and then at low levels, when cells were grown either in TB, or in NZYM in the presence of glucose, and at the lower temperature (30°C) during the induction phase. These results indicated that the bacterial growth in a rich medium, accompanied by a decrease of bacterial metabolism due to a lower temperature during protein expression induction, is an essential prerequisite for the production of the amyloidogenic L75P variant of ApoA‐I. This is not surprising since both LB and NZYM broth, in the absence of glucose, contain only a limited amount of carbohydrates and of other utilizable carbon sources. Based on these findings, TB medium and a temperature of 30°C during the induction phase were selected as the starting conditions for further optimization of the expression of the ApoA‐I proteins.

Effect of culture medium composition and temperature on recombinant ApoA‐I proteins expression levels. *Escherchia coli* cells, transformed with the vector encoding either the WT or L75P‐ApoA‐I, were grown in different media (LB, NZYM, NZYM +0.2% glucose, or TB) and protein expression was induced for 5 h following the addition of 1 mM IPTG. Protein expression was performed at both 37°C (left panels) and 30°C (right panels). Equal amounts of total bacterial cell lysates from cells grown in each culture condition were analyzed by SDS‐PAGE followed by Coomassie staining (upper panels) and Western blot analysis (lower panels). Arrows indicate ApoA‐I protein species.

Establishment of a novel system for efficient expression of L75P‐ApoA‐I

Escherichia coli cells, transformed with the vector encoding WT or L75P‐ApoA‐I (and also with vectors encoding G26R‐ApoA‐I, L174S‐ApoA‐I and L178H‐ApoA‐I, see Fig. S1), were inoculated in TB and grown in shaking flasks at 37°C for 12 h. To boost cell growth and protein production, fresh nutrients were added to the culture prior to induction of protein expression by IPTG addition. This was achieved by adding concentrated TB media (10×) to obtain a final TB concentration of four times (4×) more nutrients compared to standard TB. The potential of maltose as an additional source of carbon to improve the expression of WT and L75P‐ApoA‐I was also analyzed and compared to protein expression in the absence of maltose. As described above, higher levels of protein expression were achieved when inducing and culturing the cells at 30°C, thus this temperature was also used here for cells grown in TB 4×. However, in order to enable direct comparison with the previously used production protocol,9, 27, 28 and also to allow for comparative structural and functional analyses in relation to earlier findings on WT‐ApoA‐I,8 a temperature of 37°C was used throughout the expression in LB. The optical density (which correlates to cell number) reached at the end of the induction phase and the expression levels of WT and L75P‐ApoA‐I proteins are shown in Figure 2. As could be expected with media rich in nutrients, cell growth in TB 4× during the induction phase resulted in much higher optical density, compared to growth in LB [Fig. 2(a)]. Specifically, the bacterial culture expressing the WT protein in TB 4× reached values of optical density that were 4.3‐ to 4.7‐fold higher compared with the LB cultures. Similarly, very high optical densities were obtained for cells expressing L75P‐ApoA‐I when grown in TB 4×, indicating that expression of the amyloidogenic protein is not toxic to the cells. The TB 4× media appeared to contain sufficient carbon sources for cell growth, as the addition of maltose during the induction phase did not result in any significant differences in the optical density for the cultures expressing either WT or L75P‐ApoA‐I proteins. Estimation of the protein expression levels by SDS‐PAGE combined with Coomassie staining of total proteins and immunologic detection of the ApoA‐I species [Fig. 2(b); see also Fig. S1 for expression levels of G26R‐ApoA‐I, L174S‐ApoA‐I and L178H‐ApoA‐I variants] revealed that the expression conditions used resulted in very high amounts of produced ApoA‐I proteins. The high nutrient conditions and the reduction of the growth temperature to 30°C were particularly effective for the production of the L75P‐ApoA‐I variant, whereas extremely low amounts were obtained at the other conditions tested. Moreover, the addition of maltose had no significant effect on cell growth or on the expression levels of neither WT nor L75P‐ApoA‐I proteins. In summary, the established culture conditions contribute to the highly improved production of amyloidogenic ApoA‐I protein in two ways: by increasing the cell density (4–5‐fold) of the culture and by increasing the level of expressed protein per cell mass. The facts that the lower temperature and the high amounts of fresh nutrients enable the bacterial cells to stay longer in the growth phase, and thus have the translational machinery active longer, are likely key components to the achieved high level of expressed protein.

Induction of protein production in TB 4x greatly enhances the expression levels of L75P‐ApoA‐I variant. *Escherichia coli* cells, transformed with the vector encoding either the WT or L75P‐ApoA‐I, were grown overnight in TB at 37°C and protein expression performed at 30°C, following the addition of 1 mM IPTG, TB 4×, in the presence or absence of 0.5% maltose (M). (a) Optical density (OD_600nm) reached at the end of the induction phase (5 h) for each bacterial cell culture condition (ND, not determined). Data shown are the mean ± SEM (N = 3). (b) SDS‐PAGE analysis of equal amounts of total bacterial cell lysates from cells grown in each culture condition followed by Coomassie staining (upper panels) and Western blot analysis (lower panels). Arrows indicate ApoA‐I protein species.

High‐yield purification of L75P‐ApoA‐I

WT and L75P‐ApoA‐I proteins were expressed in bacterial cells cultured in TB 4× media, in the presence or absence of maltose, followed by purification from the bacterial cell lysates by using a two‐step purification approach, as previously described.27 The purity of the recombinant ApoA‐I proteins was evaluated by SDS‐PAGE analysis followed by Coomassie staining, and protein yield per liter of bacterial culture was determined. The SDS‐PAGE analysis showed that the purification procedure leads to apparent high levels of purity of both the His‐tagged proteins and of the final WT and L75P‐ApoA‐I not carrying the His‐tag [Fig. 3(a)]. The purity and the identity of the recombinant ApoA‐I proteins were confirmed by mass spectroscopy analysis. The purity of the recombinant proteins was estimated to be 84, 86 and 77% for WT in LB, WT in TB 4× and L75P variant, respectively [Fig. S2(a–c)]; moreover, none of the contaminants of the top 20 proteins with the highest peptide match constituted more than 1% of total protein (Fig. S3). The sequence coverages for the WT protein produced either in LB or in TB 4× and for the L75P variant in TB 4× were all 100% [Fig. S4(a–c)].

Protein expression in TB 4× leads to a high‐yield production of ApoA‐I proteins. Induced bacterial cells, grown in TB 4× in the presence of absence of 0.5% maltose (M), were harvested and lysed, and recombinant WT and L75P‐ApoA‐I proteins purified by a two‐step immobilized metal affinity chromatography. (a) 15 μl of bacterial soluble protein fraction (SF) and 3 μg of proteins eluted after each purification step (*His* and WT or *L75P* refer to ApoA‐I proteins before and after removal of the His‐tag, respectively) were analyzed by SDS‐PAGE followed by Coomassie staining. (b) The yield per liter of bacterial culture of His‐tagged WT and L75P‐ApoA‐I proteins and (c) of the proteins after the removal of the His‐tag were also evaluated (ND, not determined). Data shown are the mean ± SEM (N = 3).

Importantly, the obtained yields of WT and L75P‐ApoA‐I proteins from cells cultured in TB 4× were significantly higher compared with the yield of the WT protein expressed in LB [Fig. 3(b,c)]. Indeed, the expression of the WT protein in TB 4× lead to a 7‐fold increase in protein yield compared to the one obtained producing the WT protein in LB (the same comparison could not be made for the L75P‐ApoA‐I protein as we were unable to produce this protein in cells cultured in LB media). Of note, the yield of untagged WT‐ApoA‐I, obtained by using the method here described, is about 70% higher than the previously reported methods for the recombinant expression of native ApoA‐I, in both E. coli and Pichia pastoris. 29, 30, 31 In addition, the yields of purified G26R‐ApoA‐I, L174S‐ApoA‐I, and L178H‐ApoA‐I variants, by using the same culture conditions and purification protocol, were determined to be in the same range as the WT protein (Table 1), which indicates that the protocol can successfully be employed for amyloidogenic ApoA‐I variants in general.

Table 1.

Yield (mg/L of Bacterial Culture) of Recombinant ApoA‐I Variants Before (His‐ApoA‐I) and After (ApoA‐I) the Removal of the His‐tag

	Protein Yield (mg/L of bacterial culture)
	His‐ApoA‐I	ApoA‐I
WT	132.4 ± 20.0	74.4 ± 19.6
G26R	130.9 ± 30.4	77.3 ±12.9
L75P	81.3 ± 6.4	46.3 ± 3.7
L174S	132.7 ± 4.1	83.2 ± 6.5
L178H	146.7 ± 20.4	90.7 ± 10.5

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Data are the means ± SEM (N = 3).

Structural and functional analysis of ApoA‐I proteins produced with the high‐yield system

In order to ascertain that the new high‐yield expression and purification system had no deleterious impact on ApoA‐I structure and function, conformational structure analysis of WT and L75P‐ApoA‐I proteins was next performed followed by quantitative and qualitative analyses of their lipid binding ability.

First, the WT protein produced either in LB or in TB 4×, as well as the L75P variant produced in TB 4× were analyzed by Blue Native gel electrophoresis. As shown in Figure 4(a), all the recombinant ApoA‐I proteins presented one main protein species, indicating that the proteins are essentially monomeric at the condition tested.

Recombinant ApoA‐I proteins produced with the high‐yield system retain their structural and functional properties. (a) Blue Native gel electrophoresis of WT and L75P‐ApoA‐I proteins. The gel was loaded with 5 μg of each recombinant protein from a 0.5 mg/mL protein solution. (b) CD spectra of WT and L75P‐ApoA‐I proteins. Proteins (0.5 mg/mL) were scanned by CD spectroscopy in the far‐UV range of 200–260 nm, at 25°C. (c) Intrinsic fluorescence spectra of the recombinant ApoA‐I proteins. Intrinsic fluorescence of the ApoA‐I proteins (0.1 mg/mL) was acquired upon excitation of tyrosine and tryptophan residues at 280 nm (d) Thermal unfolding of WT and L75P‐ApoA‐I. Proteins (0.5 mg/mL) were analyzed by CD spectroscopy and the CD signal at 220 nm was recorded as a function of temperature, in the 25–95°C range. (e) Lipid Clearance Assay. Recombinant proteins (0.5 mg/mL) were incubated with DMPC multilamellar vesicles at 1:100 protein to lipid molar ratio and the decrease of the absorbance at 325 nm was measured every 10 s for 10 min. (f) Native gel electrophoresis of POPC:FC:ApoA‐I particles (rHDL) obtained by incubating the recombinant proteins with POPC and cholesterol at a 80:4:1 lipids to protein molar ratio.

Second, CD spectroscopy was used to compare the conformation of the WT protein produced either in LB or in TB 4×, as well as of the L75P variant produced in TB 4×. CD spectroscopy provides information on protein secondary structure in solution and is therefore commonly used to assay for global structural integrity.32, 33 As shown in Figure 4(b), there was no significant difference between the CD spectra obtained for the WT proteins produced in either LB or TB 4× indicating that the high‐yield expression protocol is not affecting ApoA‐I secondary structure composition. Moreover, the CD spectrum from L75P‐ApoA‐I [Fig. 4(b)] showed a lower signal intensity compared to the WT protein, denoting a lower α‐helical content, which is in agreement with previous findings.8, 24

Protein stability was next analyzed by utilizing intrinsic fluorescence of tyrosine and tryptophan residues that both emit fluorescence when excited at 280 nm. The characteristics of the emission spectra correlate to solvent exposure of these residues. Changes in the maximum emission wavelength can, therefore, be used to pinpoint protein conformational rearrangement. Thus, intrinsic fluorescence was used to highlight differences in the overall conformation of the WT protein produced either in LB or in TB 4×, and of the L75P variant produced in TB 4×. As shown in Figure 4(c), no difference was detected between the WT proteins produced in either of the culture conditions, whereas L75P intrinsic fluorescence spectra was characterized by a lower fluorescence intensity and with a maximum intensity at a longer wavelength. In agreement with previous finding, this shift is diagnostic of a more relaxed structure of the L75P‐ApoA‐I.24 The thermal stabilities of the ApoA‐I proteins were analyzed by recording the CD signal at 220 nm. As expected, there was no significant difference in the thermal denaturation curves obtained for the WT protein produced either in LB or in TB 4×. The curve obtained for the L75P‐ApoA‐I indicated an expected lower thermal stability for this variant [Fig. 4(d)], as previously shown.24 Furthermore, the thermal unfolding process was completely reversible for the WT proteins as well as for the L75P variant (Fig. S5).

A main function of the ApoA‐I protein is the binding and transport of lipids. A lipid clearance assay was therefore used to assess the ability of the ApoA‐I proteins to bind lipids. No differences in the DMPC clearance kinetics of the WT proteins produced under the different expression conditions were observed, whereas the L75P variant showed the expected slower lipid binding kinetics [Fig. 4(e)].8

Finally, the ability of the proteins to form HDL lipid‐protein complexes was analyzed and compared. Clearly, the WT and L75P‐ApoA‐I proteins produced in TB 4× media retained the ability to form rHDL particles with an estimated diameter of approximately 9.6 nm, upon incubation with POPC and cholesterol at a 1:80:4 protein to lipids molar ratio [Fig. 4(f)]. These results indicate that the high‐yield method for production of WT and amyloidogenic proteins here described results in correctly folded and functionally active ApoA‐I proteins.

Conclusion

ApoA‐I‐related amyloidosis is a rare disease for which no therapeutic approach is currently available. The availability of an efficient system for the recombinant expression of ApoA‐I amyloidogenic variants is of key importance for the development of therapeutics. However, the low stability and the tendency to aggregate that characterize these proteins make their production very challenging, which also greatly limits the possibility to perform biochemical and cellular studies aimed to unravel the molecular mechanism underlying this disease.

Here, we describe a new, highly efficient, production protocol for the recombinant expression of ApoA‐I amyloidogenic variants in E. coli. The method is based on the addition of high amounts of fresh nutrients prior the induction step and on a lower temperature during the expression phase, which combined allows for increased cell density of the bacterial culture and increased levels of expressed protein per cell mass. The new protocol enables the production of correctly folded and functional recombinant ApoA‐I variants with a yield per liter of bacterial culture, markedly higher compared with the already available protocols adapted for the recombinant production of the native ApoA‐I. The accessibility to ample amounts of recombinant ApoA‐I amyloidogenic variants will enable detailed functional and structural studies that aim to unveil the molecular mechanism underlying ApoA‐I amyloidosis, and also accelerate the development of effective therapeutic strategies.

Materials and Methods

Expression and purification of recombinant human ApoA‐I proteins

Human ApoA‐I proteins, containing a His‐tag and tobacco etch virus (TEV) protease recognition site at the N‐terminus,25 were expressed in the bacterial E. coli BL21(DE3) pLysS strain (Invitrogen, Thermo Fisher Scientific, MA) using vector pEXP‐5. Cells were cultivated in media with different formulations such as LB (10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl, Fisher Scientific, MA), NZYM (5 g/L NaCl, 2 g/L MgSO₄ 7 H₂O, 5 g/L yeast extract, 10 g/L casein hydrolysate, Sigma‐Aldrich, MO) in the presence or absence of 0.2% glucose, TB (12 g/L tryptone, 24 g/L yeast extract, 0.5% glycerol, 89 mM phosphate buffer), and TB 4× (48 g/L tryptone, 96 g/L yeast extract, 2% glycerol, 356 mM phosphate buffer) in the presence or absence of 0.5% maltose, all supplemented with 50 μg/mL ampicillin and 34 μg/mL chloramphenicol, at 30°C or 37°C. When cultures reached an OD_600nm of 0.5–0.7, protein expression was induced for 5 h following the addition of 1 mM isopropyl‐beta‐thiogalactopyranoside (IPTG, Thermo Fisher Scientific, MA). In the case of TB 4× expression medium, protein expression was induced by adding 1 mM IPTG and 10× concentrated TB, to give a 4× final concentration, to the overnight preculture. Protein expression levels were evaluated by SDS‐PAGE, followed by Coomassie staining and Western Blot analysis.

Recombinant proteins were then purified using immobilized metal affinity chromatography (His‐Trap‐Nickel‐chelating columns, GE Healthcare, IL) followed by TEV protease treatment and a second immobilized metal affinity chromatography step to remove the His‐tag.7, 27 Protein purity was analyzed by SDS‐PAGE, followed by Coomassie staining, and concentration was determined by using a NanoDrop 2000c spectrophotometer (Thermo Fisher Scientific, MA).

Western Blot analysis

Total bacterial cell lysates were separated by 4–15% Tris‐Glycine pre‐casted gels (BioRad, CA), transferred to nitrocellulose membranes, and probed with anti‐human ApoA‐I antibodies (Q0496, Dako, Agilent Technologies, CA). Detection was performed by using HRP‐conjugated secondary antibodies (GE Healthcare, IL) and a chemiluminescence detection substrate (Super‐Signal West Femto, Thermo Fisher Scientific, MA). Blots were imaged using the Odyssey Fc system (LI‐COR Biosciences, NE).

Mass spectrometric analysis

In order to estimate the purity of the recombinant proteins, the trypsin MS data from Q Exactive HFX MS (see Supplementary Materials and methods) was searched against the single‐entry protein FASTA database using Thermo Biopharma Finder v3.0. The search parameters included S/N ratio set at default value 128, trypsin, an mass accuracy set to 5 ppm with minimum confidence 0.8 to ensure low level of false‐positive identifications. The quantitative values are presented as % abundance, based on the sum of all modified components divided by the sum of all components.

The sequence coverage of the recombinant proteins was obtained searching for the pepsin digest MS data (see Supplementary Materials and methods) against the single‐entry protein FASTA database using the same software. The search parameters included S/N ratio set at default value 64, low specificity pepsin, a mass accuracy set to 5 ppm with minimum confidence 0.8 to ensure low level of false positive‐identifications. The quantitative values are presented as % abundance, based on the sum of all modified components divided by the sum of all components.

The raw DDA data were analyzed with Proteome Discoverer™ 2.2 Software (Thermo Fisher Scientific) for the determination of contaminants. Peptides were identified using SEQUEST HT against UniProtKB human database (release 20171218) and E. coli database (released 20180912). The search was performed with cysteine carbamidomethylation as static modification and N‐terminal acetylation, methionine oxidation, and phosphorylation as dynamic modifications. Precursor tolerance was set to 10 ppm and fragment tolerance was set to 0.05 ppm. Up to two missed cleavages were allowed and Percolator was used for peptide validation at a q‐value of maximum 0.01. Filter settings at the protein level were Master is equal to Master, Protein Unique Peptides is greater than or equal to 2. Filter settings peptide level were Confidence is equal to high. Twenty proteins with the highest peptide‐spectrum match (number of MS2 spectra that is associated with peptide identification for a certain protein) were used for analysis of protein contaminants.

Formation of reconstituted HDL

Lyophilized POPC (1‐palmitoyl‐2‐oleoyl‐sn‐glycero‐3‐phosphocholine, Avanti Polar Lipids, AL) and cholesterol (FC) (Avanti Polar Lipids, AL) were dissolved in 3:1 chloroform:methanol, and the solvent was evaporated by overnight incubation under a stream of nitrogen gas. POPC and FC were dissolved in PBS, and lipoparticles were generated by using the cholate dialysis method34. Briefly, POPC:FC lipoparticles were produced by incubating POPC and cholesterol, diluted in sodium deoxycholate, with ApoA‐I variants at 80:4:1 molar ratio and at a 0.5 mg/mL protein concentration. Mixtures were incubated at 37°C for 1 h and then dialyzed against PBS for 72 h. At the end of the incubation, homogeneous 9.6 nm POPC:FC:ApoA‐I particles were isolated by a size‐exclusion chromatography by using a preparative Superose 6 increase 10/300 GL column (GE Healthcare, IL). Samples were eluted in PBS, at a flow rate of 0.5 mL/min, and analyzed by Blue Native PAGE by using the NativePAGE Bis‐Tris Gel System 4–16% (Invitrogen, Thermo Fisher Scientific, MA) according to the manufacturer's instructions.

Lipid clearance assay

Lyophilized DMPC (1,2‐dimyristoyl‐sn‐glycero‐3‐phosphocholine, Avanti Polar Lipids, AL) was dissolved in 3:1 chloroform:methanol and the solvent was evaporated by overnight incubation under nitrogen gas. In order to create multilamellar vesicles, DMPC was then dissolved in PBS and the lipid suspension extruded through a 100‐nm polycarbonate membrane using the LiposoFast system (Avestin, Germany). ApoA‐I proteins were mixed with DMPC multilamellar vesicles at a 1:100 protein to lipid molar ratio and DMPC clearance was monitored by measuring the absorbance of mixture at 325 nm for 10 min, as previously described.28

CD spectroscopy

CD spectroscopy measurements were performed on a Jasco J‐810 spectropolarimeter equipped with a Jasco CDF‐426S Peltier set to 25°C. Proteins (0.5 mg/mL) were loaded into a 1 mm quartz cuvette and the spectra acquired in the far‐UV range 200–260 nm, with a 1 nm wavelength increment. Averages of five scans were baseline‐subtracted (PBS) as previously described.25

For the thermal stability analysis, CD signal at 220 nm was acquired in the 25–95°C range, with a 1°C increment.

Intrinsic fluorescence analysis

Intrinsic fluorescence emission spectra were acquired at 25°C in a 10‐mm cell by using a Jasco J‐810 spectropolarimeter equipped with a Jasco CDF‐426S Peltier and a FMO‐427S fluorescence module. Spectra were recorded in the range 300–450 nm, following excitation at 280 (tyrosine/tryptophan), with a scan speed of 150 nm/min and 5 nm slit width. Spectra are the average of five scans.

Supporting information

Figure S1. Protein production in TB 4× can be successfully used for the high‐level expression of ApoA‐I amyloidogenic variants. Escherichia coli cells, transformed with vectors encoding the ApoA‐I proteins (WT, G26R, L75P, L174S, and L178H), were grown overnight in TB at 37°C and protein expression was performed at 30°C, following the addition of 1 mM IPTG and TB 4×. Equal amounts of total bacterial cell lysates were analyzed by SDS‐PAGE, followed by Coomassie staining. Arrows indicate ApoA‐I protein species.

Figure S2. Purity of the recombinant ApoA‐I preparations. Proteins preparations (30 μg) were digested with trypsin and analyzed by LC–MS/MS. MS/MS data for (a) WT protein produced in LB (b) WT protein produced in TB 4× and (c) L75P‐ApoA‐I variant produced in TB 4×.

Figure S3. Determination the abundance of the top 20 protein species in the recombinant ApoA‐I samples. Protein samples (30 μg) were digested with trypsin and analyzed by LC–MS/MS. The peptides were identified by using UniProtKB human and E. coli databases. The abundance of each of the 20 proteins with the highest score are reported for all the three protein preparations.

Figure S4. Sequence coverage of recombinant ApoA‐I proteins. Protein samples (50 pmol) were digested with pepsin and analyzed by LC–MS/MS. Sequence coverage for (a) WT protein produced in LB, (b) WT protein produced in TB 4×, and (c) L75P‐ApoA‐I variant produced in TB 4×.

Figure S5. Thermal unfolding of the ApoA‐I proteins is a reversible process. Proteins (0.5 mg/mL) that underwent thermal unfolding were tested for their ability to refold by acquiring the CD signal at 220 nm in the 95–25°C range.

Click here for additional data file.^{(2.1MB, pdf)}

ACKNOWLEDGMENTS

This work was supported by grants from the Swedish Research Council (K2014‐54X‐22426‐01‐3 and 2009‐1039, Strategic research area Exodiab), the Vinnova – Sweden's Innovation Agency (2015‐01549), the Blanceflor foundation, the Royal Physiographic Society in Lund, the Crafoord foundation, and by the Swedish Foundation for Strategic Research (IRC15‐0067). The authors would like to thank Charlotte Welinder for excellent support with the mass spectrometry analyses and the Local MS Support at Medical Faculty, Lund University, Sweden.

Statement of importance: A highly efficient protocol for the recombinant expression of ApoA‐I amyloidogenic proteins in Escherichia coli was established. This new method allows us to obtain large amounts of these otherwise difficult‐to‐produce proteins, due to their instability and their aggregation propensity. The availability of high amounts of ApoA‐I amyloidogenic variants will enable detailed functional and structural studies, thus expanding the knowledge about the molecular basis of the disease and accelerating the development of effective therapeutic strategies.

Contributor Information

Rita Del Giudice, Email: rita.del_giudice@med.lu.se.

Jens O. Lagerstedt, Email: jens.lagerstedt@med.lu.se.

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2. Pepys MB (2006) Amyloidosis. Annu Rev Med 57:223–241. [DOI] [PubMed] [Google Scholar]
3. Langkilde AE, Herranz‐Trillo F, Bernadó PVB. Amyloid proteins In: Sigurdsson E, Calero M, Gasset M, Eds, 2018. Amyloid Proteins. Methods in Molecular Biology. New York, USA: Humana Press. Volume 1779; p. 209–239. [DOI] [PubMed] [Google Scholar]
4. Gremer L, Schölzel D, Schenk C, Reinartz E, Labahn J, Ravelli RBG, Tusche M, Lopez‐Iglesias C, Hoyer W, Heise H, Willbold D, Schröder GF (2017) Fibril structure of amyloid‐β(1–42) by cryo–electron microscopy. Science 358:116–119. [DOI] [PMC free article] [PubMed] [Google Scholar]
5. Fitzpatrick AWP, Falcon B, He S, Murzin AG, Murshudov G, Garringer HJ, Crowther RA, Ghetti B, Goedert M, Scheres SHW (2017) Cryo‐EM structures of tau filaments from Alzheimer's disease. Nature 547:185–190. [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Ruggeri FS, Benedetti F, Knowles TPJ, Lashuel HA, Sekatskii S, Dietler G (2018) Identification and nanomechanical characterization of the fundamental single‐strand protofilaments of amyloid α‐synuclein fibrils. Proc Natl Acad Sci USA 115:7230–7235. [DOI] [PMC free article] [PubMed] [Google Scholar]
7. Lagerstedt JO, Cavigiolio G, Roberts LM, Hong HS, Jin LW, FitzGerald PG, Oda MN, Voss JC (2007) Mapping the structural transition in an amyloidogenic apolipoprotein A‐I. Biochemistry 46:9693–9699. [DOI] [PMC free article] [PubMed] [Google Scholar]
8. Del Giudice R, Domingo‐Espín J, Iacobucci I, Nilsson O, Monti M, Monti DM, Lagerstedt JO (2017) Structural determinants in ApoA‐I amyloidogenic variants explain improved cholesterol metabolism despite low HDL levels. Biochim Biophys Acta Mol Basis Dis 1863:3038–3048. [DOI] [PubMed] [Google Scholar]
9. Del Giudice R, Nilsson O, Domingo‐Espín J, Lagerstedt JO (2017) Synchrotron radiation circular dichroism spectroscopy reveals structural divergences in HDL‐bound apoA‐I variants. Sci Rep 7:1–9. [DOI] [PMC free article] [PubMed] [Google Scholar]
10. Townsend D, Hughes E, Hussain R, Siligardi G, Baldock S, Madine J, Middleton DA (2017) Heparin and methionine oxidation promote the formation of apolipoprotein A‐I amyloid comprising α‐helical and β‐sheet structures. Biochemistry 56:1632–1644. [DOI] [PubMed] [Google Scholar]
11. Rye K‐A, Bursill CA, Lambert G, Tabet F, Barter PJ (2009) The metabolism and anti‐atherogenic properties of HDL. J Lipid Res 50:S195–S200. [DOI] [PMC free article] [PubMed] [Google Scholar]
12. Zannis VI, Chroni A, Krieger M (2006) Role of apoA‐I, ABCA1, LCAT, and SR‐BI in the biogenesis of HDL. J Mol Med 84:276–294. [DOI] [PubMed] [Google Scholar]
13. Domingo‐Espín J, Lindahl M, Nilsson‐Wolanin O, Cushman SW, Stenkula KG, Lagerstedt JO (2016) Dual actions of apolipoprotein A‐I on glucose‐stimulated insulin secretion and insulin‐independent peripheral tissue glucose uptake lead to increased heart and skeletal muscle glucose disposal. Diabetes 65:1838–1848. [DOI] [PubMed] [Google Scholar]
14. Stenkula KG, Lindahl M, Petrlova J, Dalla‐Riva J, Göransson O, Cushman SW, Krupinska E, Jones HA, Lagerstedt JO (2014) Single injections of apoA‐I acutely improve in vivo glucose tolerance in insulin‐resistant mice. Diabetologia 57:797–800. [DOI] [PMC free article] [PubMed] [Google Scholar]
15. The Emerging Risk Factors Collaboration (2009) Major lipids, apolipoproteins, and risk of vascular disease. J Am Med Assoc 302:1993–2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
16. Rosenson RS, Brewer HB, Ansell BJ, Barter P, Chapman MJ, Heinecke JW, Kontush A, Tall AR, Webb NR (2016) Dysfunctional HDL and atherosclerotic cardiovascular disease. Nat Rev Cardiol 13:48–60. [DOI] [PMC free article] [PubMed] [Google Scholar]
17. Siebel AL, Heywood SE, Kingwell BA (2015) HDL and glucose metabolism: current evidence and therapeutic potential. Front Pharmacol 6:258. [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Obici L, Franceschini G, Calabresi L, Giorgetti S, Stoppini M, Merlini G, Bellotti V (2006) Structure, function and amyloidogenic propensity of apolipoprotein A‐I. Amyloid 13:191–205. [DOI] [PubMed] [Google Scholar]
19. Rowczenio D, Dogan A, Theis JD, Vrana JA, Lachmann HJ, Wechalekar AD, Gilbertson JA, Hunt T, Gibbs SDJ, Sattianayagam PT, Pinney JH, Hawkins PN, Gillmore JD (2011) Amyloidogenicity and clinical phenotype associated with five novel mutations in apolipoprotein A‐I. Am J Pathol 179:1978–1987. [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Obici L, Palladini G, Giorgetti S, Bellotti V, Gregorini G, Arbustini E, Verga L, Marciano S, Donadei S, Perfetti V, Calabresi L, Bergonzi C, Scolari F, Merlini G (2004) Liver biopsy discloses a new apolipoprotein A‐I hereditary amyloidosis in several unrelated Italian families. Gastroenterology 126:1416–1422. [DOI] [PubMed] [Google Scholar]
21. Nichols WC, Gregg RE, Brewer HB, Benson MD (1990) A mutation in apolipoprotein A‐I in the Iowa type of familial amyloidotic polyneuropathy. Genomics 8:318–323. [DOI] [PubMed] [Google Scholar]
22. Mangione P (2001) Amyloid fibrils derived from the apolipoprotein A1 Leu174Ser variant contain elements of ordered helical structure. Protein Sci 10:187–199. [DOI] [PMC free article] [PubMed] [Google Scholar]
23. De Sousa MM, Vital C, Ostler D, Fernandes R, Pouget‐Abadie J, Carles D, Saraiva MJ (2000) Apolipoprotein AI and transthyretin as components of amyloid fibrils in a kindred with apoAI Leu178His amyloidosis. Am J Pathol 156:1911–1917. [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Del Giudice R, Arciello A, Itri F, Merlino A, Monti M, Buonanno M, Penco A, Canetti D, Petruk G, Monti SM, Relini A, Pucci P, Piccoli R, Monti DM (2016) Protein conformational perturbations in hereditary amyloidosis: differential impact of single point mutations in ApoAI amyloidogenic variants. Biochim Biophys Acta Gen Subj 1860:434–444. [DOI] [PubMed] [Google Scholar]
25. Petrlova J, Duong T, Cochran MC, Axelsson A, Morgelin M, Roberts LM, Lagerstedt JO (2012) The fibrillogenic L178H variant of apolipoprotein A‐I forms helical fibrils. J Lipid Res 53:390–398. [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Das M, Mei X, Jayaraman S, Atkinson D, Gursky O (2014) Amyloidogenic mutations in human apolipoprotein A‐I are not necessarily destabilizing – a common mechanism of apolipoprotein A‐I misfolding in familial amyloidosis and atherosclerosis. FEBS J 281:2525–2542. [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Petrlova J, Dalla‐Riva J, Mörgelin M, Lindahl M, Krupinska E, Stenkula KG, Voss JC, Lagerstedt JO (2014) Secondary structure changes in ApoA‐I Milano (R173C) are not accompanied by a decrease in protein stability or solubility. PLoS One 9:e96150. [DOI] [PMC free article] [PubMed] [Google Scholar]
28. Dalla‐Riva J, Lagerstedt JO, Petrlova J (2015) Structural and functional analysis of the apolipoproteinA‐I A164S variant. PLoS One 10:1–18. [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Ikon N, Shearer J, Liu J, Tran JJ, Feng S, Kamei A, Beckstead JA, Kiss RS, Weers PM, Ren G, Ryan RO (2017) A facile method for isolation of recombinant human apolipoprotein A‐I from E. coli . Protein Expr Purif 134:18–24. [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Ryabchenko AV, Kotova MV, Tverdohleb NV, Knyazev RA, Polyakov LM (2015) Production and analysis of biological properties of recombinant human apolipoprotein A‐I. Bull Exp Biol Med 160:129–133. [DOI] [PubMed] [Google Scholar]
31. Narasimhan Janakiraman V, Noubhani A, Venkataraman K, Vijayalakshmi M, Santarelli X (2016) High yield of recombinant human Apolipoprotein A‐I expressed in Pichia pastoris by using mixed‐mode chromatography. Biotechnol J 11:117–126. [DOI] [PubMed] [Google Scholar]
32. Siligardi G, Hussain R (2015) CD spectroscopy: an essential tool for quality control of protein folding. Methods Mol Biol 1261:255–276. [DOI] [PubMed] [Google Scholar]
33. Kelly SM, Price NC (2000) The use of circular dichroism in the investigation of protein structure and function. Curr Prot Pept Sci 1:349–384. [DOI] [PubMed] [Google Scholar]
34. Nichols AV, Gong EL, Blanche PJ, Forte TM (1983) Characterization of discoidal complexes of phosphatidylcholine, apolipoprotein A‐I and cholesterol by gradient gel electrophoresis. Biochim Biophys Acta Lipids Lipid Metab 750:353–364. [DOI] [PubMed] [Google Scholar]

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[pro3522-bib-0001] 1. Chiti F, Dobson CM (2017) Protein misfolding, amyloid formation, and human disease: a summary of progress over the last decade. Ann Rev Biochem 86:27–68. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0002] 2. Pepys MB (2006) Amyloidosis. Annu Rev Med 57:223–241. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0003] 3. Langkilde AE, Herranz‐Trillo F, Bernadó PVB. Amyloid proteins In: Sigurdsson E, Calero M, Gasset M, Eds, 2018. Amyloid Proteins. Methods in Molecular Biology. New York, USA: Humana Press. Volume 1779; p. 209–239. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0004] 4. Gremer L, Schölzel D, Schenk C, Reinartz E, Labahn J, Ravelli RBG, Tusche M, Lopez‐Iglesias C, Hoyer W, Heise H, Willbold D, Schröder GF (2017) Fibril structure of amyloid‐β(1–42) by cryo–electron microscopy. Science 358:116–119. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0005] 5. Fitzpatrick AWP, Falcon B, He S, Murzin AG, Murshudov G, Garringer HJ, Crowther RA, Ghetti B, Goedert M, Scheres SHW (2017) Cryo‐EM structures of tau filaments from Alzheimer's disease. Nature 547:185–190. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0006] 6. Ruggeri FS, Benedetti F, Knowles TPJ, Lashuel HA, Sekatskii S, Dietler G (2018) Identification and nanomechanical characterization of the fundamental single‐strand protofilaments of amyloid α‐synuclein fibrils. Proc Natl Acad Sci USA 115:7230–7235. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0007] 7. Lagerstedt JO, Cavigiolio G, Roberts LM, Hong HS, Jin LW, FitzGerald PG, Oda MN, Voss JC (2007) Mapping the structural transition in an amyloidogenic apolipoprotein A‐I. Biochemistry 46:9693–9699. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0008] 8. Del Giudice R, Domingo‐Espín J, Iacobucci I, Nilsson O, Monti M, Monti DM, Lagerstedt JO (2017) Structural determinants in ApoA‐I amyloidogenic variants explain improved cholesterol metabolism despite low HDL levels. Biochim Biophys Acta Mol Basis Dis 1863:3038–3048. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0009] 9. Del Giudice R, Nilsson O, Domingo‐Espín J, Lagerstedt JO (2017) Synchrotron radiation circular dichroism spectroscopy reveals structural divergences in HDL‐bound apoA‐I variants. Sci Rep 7:1–9. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0010] 10. Townsend D, Hughes E, Hussain R, Siligardi G, Baldock S, Madine J, Middleton DA (2017) Heparin and methionine oxidation promote the formation of apolipoprotein A‐I amyloid comprising α‐helical and β‐sheet structures. Biochemistry 56:1632–1644. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0011] 11. Rye K‐A, Bursill CA, Lambert G, Tabet F, Barter PJ (2009) The metabolism and anti‐atherogenic properties of HDL. J Lipid Res 50:S195–S200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0012] 12. Zannis VI, Chroni A, Krieger M (2006) Role of apoA‐I, ABCA1, LCAT, and SR‐BI in the biogenesis of HDL. J Mol Med 84:276–294. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0013] 13. Domingo‐Espín J, Lindahl M, Nilsson‐Wolanin O, Cushman SW, Stenkula KG, Lagerstedt JO (2016) Dual actions of apolipoprotein A‐I on glucose‐stimulated insulin secretion and insulin‐independent peripheral tissue glucose uptake lead to increased heart and skeletal muscle glucose disposal. Diabetes 65:1838–1848. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0014] 14. Stenkula KG, Lindahl M, Petrlova J, Dalla‐Riva J, Göransson O, Cushman SW, Krupinska E, Jones HA, Lagerstedt JO (2014) Single injections of apoA‐I acutely improve in vivo glucose tolerance in insulin‐resistant mice. Diabetologia 57:797–800. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0015] 15. The Emerging Risk Factors Collaboration (2009) Major lipids, apolipoproteins, and risk of vascular disease. J Am Med Assoc 302:1993–2000. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0016] 16. Rosenson RS, Brewer HB, Ansell BJ, Barter P, Chapman MJ, Heinecke JW, Kontush A, Tall AR, Webb NR (2016) Dysfunctional HDL and atherosclerotic cardiovascular disease. Nat Rev Cardiol 13:48–60. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0017] 17. Siebel AL, Heywood SE, Kingwell BA (2015) HDL and glucose metabolism: current evidence and therapeutic potential. Front Pharmacol 6:258. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0018] 18. Obici L, Franceschini G, Calabresi L, Giorgetti S, Stoppini M, Merlini G, Bellotti V (2006) Structure, function and amyloidogenic propensity of apolipoprotein A‐I. Amyloid 13:191–205. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0019] 19. Rowczenio D, Dogan A, Theis JD, Vrana JA, Lachmann HJ, Wechalekar AD, Gilbertson JA, Hunt T, Gibbs SDJ, Sattianayagam PT, Pinney JH, Hawkins PN, Gillmore JD (2011) Amyloidogenicity and clinical phenotype associated with five novel mutations in apolipoprotein A‐I. Am J Pathol 179:1978–1987. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0020] 20. Obici L, Palladini G, Giorgetti S, Bellotti V, Gregorini G, Arbustini E, Verga L, Marciano S, Donadei S, Perfetti V, Calabresi L, Bergonzi C, Scolari F, Merlini G (2004) Liver biopsy discloses a new apolipoprotein A‐I hereditary amyloidosis in several unrelated Italian families. Gastroenterology 126:1416–1422. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0021] 21. Nichols WC, Gregg RE, Brewer HB, Benson MD (1990) A mutation in apolipoprotein A‐I in the Iowa type of familial amyloidotic polyneuropathy. Genomics 8:318–323. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0022] 22. Mangione P (2001) Amyloid fibrils derived from the apolipoprotein A1 Leu174Ser variant contain elements of ordered helical structure. Protein Sci 10:187–199. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0023] 23. De Sousa MM, Vital C, Ostler D, Fernandes R, Pouget‐Abadie J, Carles D, Saraiva MJ (2000) Apolipoprotein AI and transthyretin as components of amyloid fibrils in a kindred with apoAI Leu178His amyloidosis. Am J Pathol 156:1911–1917. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0024] 24. Del Giudice R, Arciello A, Itri F, Merlino A, Monti M, Buonanno M, Penco A, Canetti D, Petruk G, Monti SM, Relini A, Pucci P, Piccoli R, Monti DM (2016) Protein conformational perturbations in hereditary amyloidosis: differential impact of single point mutations in ApoAI amyloidogenic variants. Biochim Biophys Acta Gen Subj 1860:434–444. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0025] 25. Petrlova J, Duong T, Cochran MC, Axelsson A, Morgelin M, Roberts LM, Lagerstedt JO (2012) The fibrillogenic L178H variant of apolipoprotein A‐I forms helical fibrils. J Lipid Res 53:390–398. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0026] 26. Das M, Mei X, Jayaraman S, Atkinson D, Gursky O (2014) Amyloidogenic mutations in human apolipoprotein A‐I are not necessarily destabilizing – a common mechanism of apolipoprotein A‐I misfolding in familial amyloidosis and atherosclerosis. FEBS J 281:2525–2542. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0027] 27. Petrlova J, Dalla‐Riva J, Mörgelin M, Lindahl M, Krupinska E, Stenkula KG, Voss JC, Lagerstedt JO (2014) Secondary structure changes in ApoA‐I Milano (R173C) are not accompanied by a decrease in protein stability or solubility. PLoS One 9:e96150. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0028] 28. Dalla‐Riva J, Lagerstedt JO, Petrlova J (2015) Structural and functional analysis of the apolipoproteinA‐I A164S variant. PLoS One 10:1–18. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0029] 29. Ikon N, Shearer J, Liu J, Tran JJ, Feng S, Kamei A, Beckstead JA, Kiss RS, Weers PM, Ren G, Ryan RO (2017) A facile method for isolation of recombinant human apolipoprotein A‐I from E. coli . Protein Expr Purif 134:18–24. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pro3522-bib-0030] 30. Ryabchenko AV, Kotova MV, Tverdohleb NV, Knyazev RA, Polyakov LM (2015) Production and analysis of biological properties of recombinant human apolipoprotein A‐I. Bull Exp Biol Med 160:129–133. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0031] 31. Narasimhan Janakiraman V, Noubhani A, Venkataraman K, Vijayalakshmi M, Santarelli X (2016) High yield of recombinant human Apolipoprotein A‐I expressed in Pichia pastoris by using mixed‐mode chromatography. Biotechnol J 11:117–126. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0032] 32. Siligardi G, Hussain R (2015) CD spectroscopy: an essential tool for quality control of protein folding. Methods Mol Biol 1261:255–276. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0033] 33. Kelly SM, Price NC (2000) The use of circular dichroism in the investigation of protein structure and function. Curr Prot Pept Sci 1:349–384. [DOI] [PubMed] [Google Scholar]

[pro3522-bib-0034] 34. Nichols AV, Gong EL, Blanche PJ, Forte TM (1983) Characterization of discoidal complexes of phosphatidylcholine, apolipoprotein A‐I and cholesterol by gradient gel electrophoresis. Biochim Biophys Acta Lipids Lipid Metab 750:353–364. [DOI] [PubMed] [Google Scholar]

PERMALINK

High‐efficient bacterial production of human ApoA‐I amyloidogenic variants

Rita Del Giudice

Jens O Lagerstedt