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Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin
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Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin Baolong Niu a,b , Bingzhang Li a,b , Huifang Wang a , Ruijie Guo a,b , Haijin Xu c , Mingqiang Qiao c,∗∗ , Wenfeng Li a,b,∗ a Key Laboratory of Interface Science and Engineering in Advanced Materials, Taiyuan University of Technology, Ministry of Education, Taiyuan, 030024, PR China b College of Materials Science and Engineering, Taiyuan University of Technology, Taiyuan, 030024, PR China c State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, Tianjin, 300071, PR China
a r t i c l e
i n f o
Article history: Received 24 May 2016 Received in revised form 19 October 2016 Accepted 20 October 2016 Available online xxx Keywords: Hydrophobin Self-assembly Cys3–Cys4 loop Rodlets Amphipathic
a b s t r a c t We used protein fusion technology to transplant the Cys3–Cys4 loop of HGFI (a class I hydrophobin from Grifola frondosa) into a nonamyloidogenic hydrophobin HFBI (a class II hydrophobin from Trichoderma reesei) and replace the corresponding amino acids between Cys3 and Cys4 in this protein to identify whether this loop renders it amyloidogenic. Water contact angle (WCA) and X-ray photoelectron spectroscopy (XPS) measurements demonstrated that the mutant protein HFBI-AR could form amphipathic membranes by self-assembling at the hydrophilic mica and hydrophobic polystyrene surfaces. This property enabled the mutant protein to alter the surface wettabilities of polystyrene and mica as well as to change the elemental composition of siliconized glass. Atomic force microscopy (AFM) measurements indicated that, unlike class I hydrophobins, no amyloid-like rodlets were observed on the mutant protein HFBI-AR coated mica surface. Moreover, the Cys3–Cys4 region could not catalyze the mutant protein HFBI-AR to drive intermolecular association and formation of a cross- rodlet structure to resist depolymerization in organic solvents when it selfassembled at water–air interfaces. These results demonstrate that the Cys3–Cys4 loop is not the major determinant that initiates HGFI to form rodlets or account for the unique properties of the proteins. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Hydrophobins belong to a class of small surface active proteins that are uniquely secreted by filamentous fungi [1–5]. These proteins can self-assemble into stable amphipathic films at hydrophobic–hydrophilic interfaces and are responsible for a wide variety of functions in fungal physiology [6–11]. Aside from having a characteristic pattern of eight cysteine residues (including two pairs of adjacent cysteines) and four intramolecular disulfide bonds, hydrophobins share little amino acid sequence homology [12–14]. All of these proteins can self-assemble at hydrophilic/hydrophobic
∗ Corresponding author at: Key Laboratory of Interface Science and Engineering in Advanced Materials, College of Materials Science and Engineering, Taiyuan University of Technology, Taiyuan, 030024, PR China. ∗∗ Corresponding author at: State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, No. 94 Weijin Road, Tianjin 300071, PR China. E-mail addresses: [email protected] (M. Qiao), [email protected] (W. Li).
interfaces into amphipathic films and improve surface wettability [8,15–18]. Based on the stability of the formed amphipathic monolayers and spacing of the cysteine residues, hydrophobins are categorized into two classes, I and II [19–22]. The membranes formed by class I hydrophobins are extremely robust and consist of amyloid-like fibrillar structures, known as rodlets, which can only depolymerize in strong acids, such as trifluoroacetic acid (TFA) [23–28]. By contrast, the films self-assembled by class II hydrophobins are not fibrillar and can be easily dissociated into detergent and alcohol solutions [29–31]. Based on the amino acid sequence comparison of class I and II hydrophobins, the most remarkable difference in the structures between class I and II hydrophobins is the presence of a disordered loop, such as Cys3–Cys4 and Cys7–Cys8 loops (polypeptide segments from Cys3 to Cys4 and Cys7 to Cys8, respectively) [32,33]. In the primary structure, class I hydrophobins have a much larger Cys3–Cys4 loop and the polypeptide segment between this loop displays much greater sequence variation (from 4 to 44 residues). In particular, the length of the inter-cysteine regions is highly variable. By contrast, class II hydrophobins display substantially
http://dx.doi.org/10.1016/j.colsurfb.2016.10.048 0927-7765/© 2016 Elsevier B.V. All rights reserved.
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more sequence similarity to one another and the lengths of the polypeptide segments between cysteines 3 and 4 (the Cys3–Cys4 loop) are fully conserved [13,15,24]. Protease digestion and mass spectrometry as well as hydrogen/deuterium exchange studies of class I hydrophobin SC3 show that the Cys3–Cys4 loop has the propensity to form an ␣-helix structure when it initially adheres to hydrophilic/hydrophobic interfaces [34]. Moreover, the molecular dynamics simulations of SC3 show that the Cys3–Cys4 loop is a major driver when SC3 self-assembles into rodlet structures [34,35]. However, when the number of amino acid residues in the Cys3-Cys4 loop was reduced to 15 residues in class I EAS hydrophobin, rodlet formation was not impaired and the surface activity and properties of rodlets were indistinguishable from the full length protein [13]. HGFI is a class I hydrophobin that is secreted by the edible mushroom Grifola frondosa. In a previous study, we found that HGFI could form typical rodlet structures on the mica surface [36]. The membranes formed by HGFI were robust and could not depolymerize in a hot detergent and alcohol solution [37,38]. Moreover, circular dichroism (CD) shows that self-assembly of HGFI at the water–air interface could give rise to significant secondary structural changes [38], which indicates that the interfacial assembly yields highly ordered -sheets to promote rodlet formation. To study the role of the Cys3–Cys4 loop in class I hydrophobin, we previously obtained a mutant protein HGFI-AR by replacing the Cys3–Cys4 loop of HGFI with the loops that are present in typical class II hydrophobin HFBI using protein fusion technology. The results show that the rodlet structures of HGFI are impaired and the membranes formed by the mutant protein become less robust after the Cys3–Cys4 loop is replaced [39]. Therefore, we speculate that the Cys3–Cys4 loop may play by major determinant in the formation of the rodlet structure of HGFI. To further investigate the contribution of the Cys3–Cys4 loop to the rodlet structure assembly and function of HGFI, we use protein fusion technology to transplant the Cys3–Cys4 region of HGFI into a nonamyloidogenic hydrophobin HFBI to identify whether this loop can render it amyloidogenic and change its physicochemical properties. 2. Materials and methods 2.1. Strains, vectors and reagents Pichia pastoris (P. pastoris) GS115, Escherichia coli (E. coli) DH5␣ and plasmid pPIC9 were purchased from Invitrogen (Beijing, China). pET-28a-hgfI and pMD19-T-hfbI were obtained from Prof. M.Q. Qiao (State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, China). All of the restriction enzymes, polymerases, and ligases were purchased from TaKaRa (Dalian, China). DNA purification kits, DNA and protein markers were purchased from TaKaRa (Dalian, China). All of the primers used in this study were synthesized by Takara. Other reagents and chemicals were purchased from Sigma or TaKaRa.
Table 1 Primers used in the current study. Name
Sequence 5 –3
P1 P2 P3 P4 P5 P6 5 AOX 3 AOX
CGCTCGAGAAAAGAAGCAACGGCAACGGCAATG AGTGGAGGTAGACTCACAGCACTGGGGGTT CTCGTCGGTCTCACCTGCAAAGTCCCCTCC CGGAATTCAAGCACCGACGGCGGTCTG AACCCCCAGTGCTGTGAGTCTACCTCCACT GGAGGGGACTTTGCAGGTGAGACCGACGAG GACTGGTTCCAATTGACAAGC GCAAATGGCATTCTGACATCC
The restriction site XhoI in P1 and EcoRI in P4 are in underline; the Kex2 protease cleavage site is in bold.
as primers to obtain gene fragment A3. Fourth, the A1, A2 and A3 genes were used together as templates and P1/P4 as forward and reverse primers to generate the required mutant gene hfbI-ar by PCR. Then, the obtained hfbI-ar coding sequence was subcloned into the pMD19-T vector to give pMD19-T-hfbI-ar. The correct gene was confirmed by restriction enzyme digestion and DNA sequencing and was then cloned into the pPIC9 expression vector using XhoI and EcoRI restriction enzymes to obtain a recombinant vector, pPIC9-hfbI-ar. In the recombinant pPIC9-hfbI-ar vector, the hfbI-ar gene was located downstream of the AOX1 promoter, which can be induced using methanol as a sole carbon source. To ensure secretion of HFBI-AR, the ␣-factor secretion signal was placed upstream of the expressing gene, and the final recombinant vector pPIC9-hfbI-ar was verified by the XhoI and EcoRI cleavage and gene sequence analysis using the primer pairs 5 AOX/3 AOX. The hfbI-ar gene was transformed, and mutant protein HFBIAR was expressed and purified as described in Niu et al. [40]. The plasmid pPIC9-hfbI-ar was linearized with StuI and transfected into P. pastoris GS115 (His− ) cells by electroporation using a Bio-Rad gene pulser apparatus (25 F, 200 , and 2.0 kV) according to the manufacturer s instructions. Transformants were plated on minimal dextrose (MD) medium to screen for the His+ clones. Fifty His+ clones were selected and subsequently patched on MD and minimal methanol (MM) plates to determine the His+ Mut+ phenotypes. Colony PCR and sequencing were performed using the 5 AOX1 and 3 AOX1 primers to confirm the integration of hfbI-ar into the P. pastoris genome. The HFBI-AR expression levels from fifteen fastgrowing strains (His+ Mut+ ) were analyzed by RP-HPLC after they were induced by methanol supplementation to a final concentration of 0.5% at 24 h intervals for 5 days. The clone that expressed the highest level of HFBI-AR was chosen for large-scale production. A two-step process was used to purify the HFBI-AR protein. The first step was ultrafiltration using a hollow fiber membrane module with a 4-kDa molecular weight cut off and then lyophilized. RP-HPLC was used to further purify HFBI-AR with a Vydac C4 reversed-phase column. Introduction of the correct mutation HFBIAR was identified by 16% Tricine-SDS-PAGE, Western blotting and mass spectroscopy. 2.3. Water contact angle measurements
2.2. Production of mutant protein HFBI-AR The gene hfbI-ar was obtained by PCR using a four-step procedure, and the primers used are shown in Table 1. First, the pMD19-T-hfbI vector was used as a template and P1/P2 as forward and reverse primers to obtain gene A1. The KEX2 protease cleavage site following the XhoI restriction site was inserted at the 5 -terminus of P1. Second, pMD19-T-hfbI was also used as the template and P3/P4 as forward and reverse primers to obtain gene A2. An EcoRI restriction site sequence was added to the 5 end of P4. Third, the vector pMD18-T-hgfI was used as a template and P5/P6
The cleaned polystyrene and mica were incubated overnight in a 0.02 mg/mL HFBI-AR solution at 20 ◦ C. Then, the excess solution on the modified substrates was blown off with nitrogen gas. Finally, the protein coated surfaces were gently washed using water and dried with nitrogen gas again. To identify the stability of the film formed by HFBI-AR, the modified polystyrene was also washed using a 60% (v/v) ethanol solution or 2% boiling SDS solution. The WCAs of the modified and unmodified surfaces were measured with a 5-L water droplet using an optical contact angle meter (DSA100, Krus GmbH, Hamburg, Germany) at room temperature.
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The average WCA values were obtained from three measurements at different locations. 2.4. X-ray photoelectron spectroscopy (XPS) measurements The elemental compositions of bare and HFBI-AR (100 g/mL)coated siliconized glasses were analyzed with an XPS (Kratos Axis Ultra DLD) spectrometer employing a monochromated Al–Ka X-ray radiation source (h = 1486.6 eV), hybrid (magnetic/electrostatic) optics and a multi-channel plate and delay line detector (DLD). All of the XPS spectra were recorded using an aperture slot of 300 × 700 m. The survey spectra were recorded with a pass energy of 160 eV and high resolution spectra with a pass energy of 40 eV. 2.5. Atomic force microscope (AFM) measurements A 10-L aliquot of a 0.02 mg/mL HFBI-AR solution was deposited on a freshly cleaved mica substrate, and water was allowed to evaporate overnight in a covered container. The surface morphology of the HFBI-AR membrane was obtained using a NanoScope IIIa Multimode atomic force microscope (Veeco Instruments, NY, USA) in tapping mode at ambient atmosphere. The probe used contained silicon nitride cantilevers with a nominal force constant of 50 N/m. The scan rates were approximately 1 Hz. The damping ratio (set-point amplitude/free amplitude) was typically approximately 0.7–0.8. Analysis of the images was performed using a Scanning Probe Image Processor (SPIP; Image Metrology, Lyngby, Denmark).
Fig. 1. (a) Silver-stained Tricine-SDS–PAGE of purified HFBI-AR expressed in P. pastoris. M, molecular weight standards. Lane 1, fermentation supernatant of P. pastoris transformed with the pPIC9 vector. Lane 2, the lyophilisate of HFBI-AR concentrated/buffer exchanged by ultrafiltration. Lane 3, HFBI-AR purified by RP-HPLC. (b) Western blot analysis of HFBI-AR using a polyclonal antibody against HFBI. Lane 1, fermentation supernatant of P. pastoris transformed with the pPIC9 vector. Lane 2, the purified HFBI-AR.
CR and protein were incubated at room temperature for 30 min and shaken for 5 min prior to spectral analysis. Both CR and class I hydrophobin rHGFI were used as controls. 3. Results
2.6. Circular dichroism dpectropolarimetry 3.1. Expression and purification of HFBI-AR The secondary structure changes of HFBI-AR were measured by CD after it was self-assembled at an air-water interface. HFBIAR samples at a concentration of 50 g/mL in MQW were used in this study. The CD spectra of the assembled hydrophobins were obtained after vigorously shaking on a vortex mixer for 5 min. The spectra were recorded over the wavelength range from 190 to 250 nm on a Jasco J-715 CD spectrometer (Japan) using a 1-mm quartz cuvette. The temperature was kept at 25 ◦ C, and the sample compartment was continuously flushed with nitrogen gas. The samples’ spectra were the result of averaging 5 scans obtained using 5-s averaging per point, a bandwidth of 1 nm and a stepwidth of 0.5 nm. The spectrum of the reference solution without protein was used to correct for the background signal. 2.7. Thioflavin T (ThT) staining Purified HFBI-AR was dissolved in MQW and mixed with a stock ThT solution prepared in MQW. The effective concentrations of HFBI-AR and ThT in the mixture were 40 g/mL and 5 M, respectively. The sample and control (5 M ThT aqueous solution) for fluorimetry were performed using a Cary Eclipse Fluorescence Spectrophotometer (Varian Optical Spectroscopy Instruments, Mulgrave, Australia) before and after vortexing for 5 min. An emission scan from 450 to 600 nm was performed with an excitation wavelength of 435 nm. 2.8. Congo red (CR) binding assays CR binding was monitored by measuring the absorption spectra from 300 to 700 nm using a BIO-TEK Microplate Reader model (BioTek U.S., Winooski, VT, USA). Phosphate buffer (pH 4.0) was used to blank the instrument. Phosphate buffer (50 mM, pH 4.0) was used to dissolve CR and HFBI-AR. In the sample, the final concentrations of the protein and CR were 100 g/mL and 10 g/mL CR, respectively. Mixtures of
Gene A1 (86 bp) and A2 (159 bp) fragments were amplified by PCR using the pMD19-T-hfbI vector as a template. Gene A3 (126 bp) fragments were amplified by PCR using the pET-28a-hgfI vector as a template. A 311-bp HFBI-AR coding sequence was obtained using the A1, A2 and A3 gene sequences together as the template, and it was cloned into the P. pastoris expression vector pPIC9 to construct pPIC9-hfbI-ar. The hfbI-ar gene was located downstream of the AOX1 promoter, which means that HFBI-AR production can be induced by using methanol as the sole carbon source [41]. To ensure that full-length HFBI-AR was secreted to the extracellular environment, we designed a Lys-Arg linker to add after the XhoI site, which was recognized and specifically cleaved by the yeast KEX2 protease. The constructed plasmid pPIC9-hfbI-ar was linearized with StuI and transformed into the GS115 (His− ) strains using a Gene Pulser (Bio-Rad) apparatus (25 F, 200 X, 2.0 kV), as described in the manufacturer’s instructions. Fast growth of the selected transformants (His+ Mut+ ) was continued, as described by Niu et al. [40]. After being selected on MD and MM media plates, fifteen fast-growing transformants (His+ Mut+ ) were identified by PCR and sequencing using the 5 AOX/3 AOX primers. The clone with the highest HFBI-AR production (from the identified strains) was selected after the supernatants of the fermentation medium were analyzed by RP-HPLC. Ultrafiltration was used for the first step to remove salt and small impurities as well as to concentrate the extract. RP-HPLC was used for further purification. Both the lyophilized HFBI-AR after ultrafiltration and RP-HPLC purification were analyzed by 16% Tricine-SDS-PAGE. A clear band of approximately 9 kDa (coinciding with the expected size of HFBI-AR) was found in the selected transformant, but was absent from the control strains (P. pastoris transformed with pPIC9 vector) (Fig. 1a). Purified HFBI-AR was also identified by Western blot and mass spectrometry. The results showed that HFBI-AR could specifically bind to the polyclonal anti-HFBI antibody (Fig. 1b), and the molecular weight of this protein was 9599.19 Da, which is consistent with
Please cite this article in press as: B. Niu, et al., Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin, Colloids Surf. B: Biointerfaces (2016), http://dx.doi.org/10.1016/j.colsurfb.2016.10.048
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Table 2 The WCAa on the bare or HFBI-AR modified Polystyrene and mica surfaces.
Polystyrene Rinsed with 60% ethanol Rinsed with 2% SDS Mica
Bare surface (deg)
HFBI-AR(deg)
85 ± 2
36 ± 3 82 ± 2 84 ± 3 16 ± 2
7±2
The WCA values were averaged from three measurements at different locations. a n = 3.
the calculated value (9599.39 Da). As a result, we speculated that the target protein HFBI-AR was successfully expressed in P. pastoris and correctly secreted to extracellular medium without extra N-terminal amino acids. 3.2. Membrane formation and surface activity of the HFBI-AR protein Hydrophobins have a remarkable biophysical property, wherein they can self-assemble into amphipathic films at hydrophobic–hydrophilic interfaces to reverse their wettability. To detect whether the mutant protein HFBI-AR could convert the surface from hydrophilic to hydrophobic and vice versa, surface properties were probed with WCA measurements before and after HFBI-AR modification. The values of the WCA are shown in Table 2. The WCA of the bare polystyrene surface was 85 ± 2◦ , and this value dramatically decreased to 36 ± 3◦ with HFBI-AR coating on its surface. By contrast, the WCA results of the mica surface modified by HFBI-AR showed only a slight increase from 7 ± 2◦ to 16 ± 2◦ . Remarkably, the above results demonstrate that HFBI-AR is a surface active protein that contains hydrophobic and hydrophilic patches. As a result, once this protein coats the hydrophobic polystyrene surface, the hydrophobic region can adsorb onto its surface in response to hydrophobic interactions, which exposes the hydrophilic region to the outside to renders the polystyrene surface hydrophilic. In turn, when HFBI-AR self-assembles on the mica surface, the hydrophilic region faces the hydrophilic surface, leaving the hydrophobic region exposed to the outside, which subsequently made the mica surface hydrophobic. The WCA value of the mica surface only slightly increased after HFBI-AR-modified treatment, which could be explained as follows. First, as the reviewer suggested, the hydrophobin HFBI-AR might only partially cover the support surfaces; Second, once the mica surface is coated, the hydrophilic part of the first layer faces the surface and the hydrophobic patch of the first layer turns outside; then, the hydrophobic group of the second layer binds to the hydrophobic part of the first layer; finally, the hydrophilic part points outside. To further study the stability of the film formed by HFBI-AR, the HFBI-AR- modified polystyrene surfaces were thoroughly washed with 60% ethanol or 2% hot SDS. The WCA values dramatically increased from 36 ± 3◦ to 82 ± 2◦ and 84 ± 3◦ , which are close to that of the bare polystyrene surface. The restoration of the WCA values suggested that, unlike class I hydrophobins, the membrane formed by HFBI-AR was not as robust as that easily dissociated in 60% ethanol or 2% hot SDS. To explore the self-assembling properties of HFBI-AR, we also used XPS and AFM to detect the membrane formed by this protein on different surfaces. The XPS survey spectra and relative elemental composition of the siliconized glass surface before and after HFBIAR modification are shown in Fig. 2 and Table 3, respectively. A high intensity of O 1s and Si 2p signals and low intensity of the C 1s signal were detected on siliconized glass before HFBI-AR modification, and almost no S 2p s and N1 signals were observed in the spectra of the bare siliconized glass. After the surface was coated with HFBIAR, the elemental composition of Si 2p dramatically decreased from
Fig. 2. XPS spectra of siliconized glass surfaces before and after HFBI-AR modification.
Table 3 The relative elemental compositions of the bare and HFBI-AR modified siliconized glass surfaces. Spectra sample
C1s (%)
N1s (%)
O1s (%)
Si 2p (%)
S 2p (%)
Siliconized glass Siliconized glass + HFBI-AR
8.96 59.96
0.90 14.55
55.34 22.42
22.50 2.23
0.00 0.82
22.50% to 2.23% and the C 1s and N 1s signals became predominant in the spectra. In addition, a weak intensity of S 2p was also detected. Obviously, there is no Si element in HFBI-AR, which is only presents in siliconized glass. The high-intensity of the Si 2p and O 1s peaks represents a characteristic XPS spectrum of the siliconized glass surface. However, N and S elements only belong to the HFBI-AR protein family. Therefore, the increase in the N 1s and S 2p signals and decrease in the Si 2p signal confirm that HFBI-AR can form a membrane to cover the siliconized glass surface to a greater extent. Similar changes in peak intensity also occurred in the XPS spectra before and after hydrophobin HFBI was coated on the hydrophilic mica surfaces [42], which suggested that hydrophobin could also form a membrane on the hydrophilic mica surface, leading to an intensity decrease of the Si 2s and Si 2p peaks and a signal increase of the N 1s peak. However, when we rinsed HFBI-AR-coated siliconized glass with 2% hot SDS, the elemental composition reverted back to bare siliconized glass (data not shown). The result is in accordance with the conclusion obtained from the WCA measurements, which indicates that the membrane formed by HFBI-AR is not so robust that we cannot resist washing with hot SDS and ethanol. As already mentioned, the class I (not class II) hydrophobins present with rodlet structures when their dilute solution dries on a solid surface. AFM was also used in this study to observe the topography image of HFBI-AR. As stated in the Materials and methods section, when the HFBI-AR solution droplet dried on a freshly mica surface (Fig. 3a), only a compact monolayer of protein was clearly present and there was no rodlet structure observed. This membrane is different from the results found in our previous research that showed that typical mosaic rodlet structures (approximately 100 nm in length) were formed when a recombinant class I hydrophobin HGFI (rHGFI) solution was dried on a freshly cleaved mica surface (Fig. 3b). However, its topography image in AFM is
Please cite this article in press as: B. Niu, et al., Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin, Colloids Surf. B: Biointerfaces (2016), http://dx.doi.org/10.1016/j.colsurfb.2016.10.048
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Fig. 3. AFM images of different proteins dried onto mica; the image size is 1 × 1 m and the height scale is 10 nm for (a) HFBI-AR and (b) rHGFI.
Fig. 4. CD spectra of the mutant protein HFBI-AR in water (thick line) and after vigorous shaking (dotted line). The CD spectra were the averages of 5 scans.
similar to the membrane formed by recombinant class II hydrophobin HFBI (rHFBI) [38]. 3.3. Secondary structure changes upon self-assembly of HFBI-AR and rHGFI CD spectroscopy was used to study the secondary structural changes of HFBI-AR before and after self-assembly at the air-water interface. The spectra of monomeric HFBI-AR in water showed a maximum ellipticity that was close to 190 nm and a minimum that was close to 205 nm (Fig. 4 thick line). After vigorous shaking, only a slight decrease in the intensity of the spectrum was observed, which was due to the accumulation of protein at the liquid-air interface (Fig. 4 dashed line). Contrary to HFBI-AR, we previously observed that when the class I hydrophobin rHGFI solution was vortexed for 5 min, the assemblages of rHGFI acquired a predominately -sheet structure, which caused the minimum ellipticity to shift from 205 to 215–217 nm [39]. We speculate that the lack of stacked -sheets in solution-assembled HFBI-AR is present in the interfacially assembled rHGFI points to different assembly pathways between the two proteins. It has been reported that the aromatic dye ThT could specifically bind to stacked -sheet structures in amyloid fibrils to greatly
increase its fluorescence yield [43–45]. For this reason, it has been widely used to monitor amyloid formation [46]. The rodlet structure of class I hydrophobins shares many similarities with amyloid fibers, and the rodlets from SC3, EAS and rHGFI have been shown to bind ThT [13,38,47,48]. As a result, to examine whether the mutant protein HFBI-AR had the ability to self-assemble into amyloid-like structures, such as class I hydrophobins, the fluorescence probe ThT was utilized as described in the Materials and methods section. The fluorescence intensity was observed at 485 nm in the presence of a ThT and HFBI-AR-ThT mixed solution before and after vigorous vortexing (Fig. 5a), which could drive the intermolecular association and induce the protein solution to form rodlets by maximizing the air-water interfaces. Before vortexing, the fluorescence intensity value of the ThT/HFBI-AR mixed sample was 2-fold higher than that of a pure ThT water solution, which suggests that like other hydrophobins, HFBI-AR might contain some stacked -sheets in the monomeric structure or that the protein in the solution was not completely monomeric before testing. However, an increased fluorescence intensity of the ThT/HFBI-AR solution was hardly observed after vortexing for 5 min or longer (data not shown), which indicated that no substantial -sheet structure level formed when HFBI-AR selfassembled at the air-water interface. This phenomenon is different from our previously observed results that showed that when the recombinant protein rHGFI/ThT mixed solution was vortexed for 5 min (Fig. 5b), a pronounced increase in the fluorescence intensity was detected [38,39]. Moreover, -sheet structures dramatically increased after rHGFI self-assembled at the water-air interface. As a result, we speculate that the assembly pathways for rHGFI and HFBI-AR at the air-water interface were different. As in ThT, CR is a commonly used dye for amyloid detection because it can also bind to ordered stacks of -sheets and exhibits a significant shift in its absorption spectrum [49]. Because both CR and hydrophobins are negatively charged at neutral pH, the assay was performed at pH 4.0 to avoid electrostatic repulsion and prevent binding. The spectra of protein/CR and CR alone are shown in Fig. 6. Due to the limited extent of -sheets in BSA, this protein was used as a negative control. Meanwhile, the class I hydrophobin rHGFI, which could form rodlets, was also used as a positive control. Fig. 6 shows that the spectrum shifts from 490 nm to 530 nm when the rHGFI/CR mixed solution was vigorously vortexed. Like the negative control, a structural change at the air-water interface could not be observed from HFBI-AR, which indicates that the assemblages of this protein formation are not rodlets or sufficiently unstable to cause a spectrum shift.
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4. Discussion
Fig. 5. Fluorescence emission spectra of thioflavin T in the presence of HFBI-AR or control rHGFI. (a) The sample contained ThT alone (dashed line) or was unvortexed (dotted line) or vortexed (solid line) HFBI-AR. (b) The sample contained ThT alone (dashed line) or was unvortexed (dotted line) or vortexed (solid line) rHGFI.
Fig. 6. The adsorption spectra of different Congo red solutions.
Hydrophobins are small surface active proteins. Aside from a characteristic pattern of eight cysteine residues, there is little amino acid sequence similarity between these proteins. One of the major differences between class I and II hydrophobins is that in the primary structure, class I hydrophobins have a larger variable Cys3–Cys4 loop [2,50], which is the least conserved region in the protein. However, the corresponding loop in class II hydrophobins is smaller and conserved (11 residues). The secondary structure prediction suggests that part of the Cys3–Cys4 loop in class I hydrophobin has a high tendency to form a helix (as an anchor) to strongly bind to hydrophobic surfaces [51]. This prediction was confirmed by Wang; when their research team used various mass spectrometry techniques to monitor the structural changes of SC3, they found that the Cys3–Cys4 loop had a high affinity for Teflon and accompanied the formation of the ␣-helical structure to resist dissolution, which indicates that this loop is the key region driving intermolecular association and rodlet structure formation when SC3 self-assembles at hydrophilic/hydrophobic interfaces [34]. When class II hydrophobins self-assembled on the hydrophobic surface, the function of the Cys3–Cys4 loop differed and the smaller part bound much less at the surface [52]. Kwan et al. proposed a possible model for EAS rodlet formation [25], which suggests that the Cys3–Cys4 and Cys7–Cys8 disordered loop might “add on” to the -barrels to form an additional  sheet by H-bonds at air-water interfaces. However, when they removed 17 amino acid residues from 25 of the Cys3–Cys4 loops in EAS, the core structure of the mutant EAS was not essentially impaired and rodlets were still generated, which suggested that the Cys3–Cys4 loop is not central to EAS rodlet formation and the folding structure of the EAS monomer [13,53]. We previously replaced the amino acids between Cys3 and Cys4 of the class I hydrophobin HGFI from Grifola frondosa with those between Cys3 and Cys4 of the class II hydrophobin HFBI from Trichoderma reesei. Because replacement of the Cys3–Cys4 loop could impair the rodlet formation of HGFI [39], we conclude that the Cys3–Cys4 loop in HGFI may play an important role in initiating rodlet formation. To investigate the requirement for this loop in more detail, we inserted the Cys3–Cys4 region of HGFI into HFBI (a typical nonamyloidogenic hydrophobin) to identify whether this loop can render it amyloidogenic and change its physicochemical properties. Remarkably, the physicochemical properties of the mutant protein HFBI-AR are distinguishable from the class I hydrophobin rHGFI and other reported class I hydrophobins. Transplantation the Cys3–Cys4 loop is unable to trigger intermolecular associations of HFBI-AR along with stacked -sheet generation and render rodlet formation of this protein, as judged by ThT binding and AFM. Furthermore, compared to rHGFI, the membrane formed by the mutant protein HFBI-AR could not resist 60% ethanol and 2% hot SDS solubilization. This finding is consistent with the study by Kwan et al. [13], who found that deletion of 15 residues from the Cys3–Cys4 loop in EAS does not affect the physicochemical properties of its rodlet formation or change its surface activity. The previous reports showed that despite the very low level of sequence conservation, class I and II hydrophobins shared very similar folds. The three-dimensional structure showed that both class I and II hydrophobins had almost identical -barrel folds that consisted of four antiparallel -strands. This unique fold led the hydrophobins to form an amphipathic core structure [32,52]. Although the barrel portion of the class I hydrophobin fold was very similar to that of class II hydrophobin, the remainder of the structures were different. In class I hydrophobins, two disordered loops formed by the amino acid residues between Cys3–Cys4 and Cys7–Cys8 were present, but they were absent in class II
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hydrophobins. Moreover, the results of the triple-resonance NMR test showed that a two stranded -sheet in class I hydrophobins (EAS) to the outside of the -barrel was linked by the last disulfide, but the crystal structures of the class II hydrophobins (HFBI and HFBII) as solved by X-ray crystallography showed that in class II hydrophobins, the last disulfide links to an ␣-helix [13,24,52]. According to the possible model for class I hydrophobin rodlet formation, the two disordered loops could add on to the -barrels to form an additional  sheet at air-water interfaces, leading to class I hydrophobin rodlet structure formation [32]. As a result, the interaction between the Cys3–Cys4 and Cys7–Cys8 loops at airwater interfaces might play a central role in class I hydrophobin rodlet formation. Although part of the residues from the Cys3-Cys4 loop in class I hydrophobin EAS was deleted, rodlet formation was not hampered [13]. Site-directed mutagenesis and peptide experiments showed that inserting the Cys7-Cys8 loop of EAS into a nonamyloidogenic class II hydrophobin could enable it to form rodlets, is central to class I hydrophobin rodlet formation [53]. From the above results, we speculate that the major determinant that distinguishes the surface activity and self-assembly properties of class I and II hydrophobins is the amphipathic core region instead of the Cys3–Cys4 loop. Because the Cys3–Cys4 loop is not involved in the formation of this core region of class I hydrophobins, this flexible loop may only participate in the interaction of HGFI monomers with the interface, but is not responsible for triggering polymeric structure formation. As a result, when HFBI obtains the Cys3–Cys4 loop from HGFI, the physicochemical properties and structure are essentially unchanged. This observation also indirectly demonstrates why class I hydrophobins, which have this type of a non-conserved region in both the length and amino acid composition, can also display the same fold as well as polymeric rodlet structural similarities in morphology and dimensions. 5. Conclusions To investigate the contribution of the Cys3–Cys4 loop to the rodlet structure assembly and function of class I hydrophobins, we used protein fusion technology and obtained a mutant protein HFBI-AR by replacing the amino acids between Cys3 and Cys4 of class II hydrophobin HFBI with those that were between Cys3 and Cys4 of class I hydrophobin HGFI. Like native hydrophobins, HFBIAR retained amphiphilic properties, which could lead it to form amphipathic membranes at different surfaces and change their wettabilities. However, compared to class I hydrophobins, HFBI-AR could not self-assemble into rodlets and there was an accompanying increase in the -sheet conformational change in the secondary structure, which led to formation of a film in response to this protein becoming less stable. As a result, we speculated that the Cys3–Cys4 loop in class I hydrophobin HGFI is not the major determinant that initiates HGFI to form rodlets and distinguishes the surface activity and self-assembly properties of HGFI and HFBI. Conflict of interest The authors declare that they have no conflict of interest. Acknowledgments This research was financially supported by the National Natural Science Foundation of China (31170066) and Natural Science Foundation of Shanxi Province (2014021020-3, 2015021080), Scientific and Technological Innovation Programs of Higher Education Institutions in Shanxi (2015130), and Research Project Supported by Shanxi Scholarship Council of China (2015-033).
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References [1] M.L. de Vocht, I. Reviakine, H.A.B. Wösten, A. Brisson, J.G.H. Wessels, G.T. Robillard, Structural and functional role of the disulfide bridges in the hydrophobin SC3, J. Biol. Chem. 275 (2000) 28428–28432. [2] J. Hakanpää, A. Paananen, S. Askolin, T. Nakari-Setälä, T. Parkkinen, M. Penttilä, M.B. Linder, J. Rouvinen, Atomic resolution structure of the HFBII hydrophobin, a self-assembling amphiphile, J. Biol. Chem. 279 (2004) 534–539. [3] J. Wessels, O. De Vries, S.A. Asgeirsdottir, F. Schuren, Hydrophobin genes involved in formation of aerial hyphae and fruit bodies in schizophyllum, Plant Cell 3 (1991) 793–799. [4] V.K. Morris, Q. Ren, I. Macindoe, A.H. Kwan, N. Byrne, M. Sunde, Recruitment of class I hydrophobins to the air: water interface initiates a multi-step process of functional amyloid formation, J. Biol. Chem. 286 (2011) 9. [5] J.M. Kallio, M.B. Linder, J. Rouvinen, Crystal structures of hydrophobin HFBII in the presence of detergent implicate the formation of fibrils and monolayer films, J. Biol. Chem. 282 (2007) 28733–28739. [6] R.E. Beever, G.P. Dempsey, Function of rodlets on the surface of fungal spores, Nature 272 (1978) 608–610. [7] S.O. Lumsdon, J. Green, B. Stieglitz, Adsorption of hydrophobin proteins at hydrophobic and hydrophilic interfaces, Colloid Surf. B-Biointerfaces 44 (2005) 172–178. [8] E.S. Basheva, P.A. Kralchevsky, K.D. Danov, S.D. Stoyanov, T.B.J. Blijdenstein, E.G. Pelan, A. Lips, Self-assembled bilayers from the protein HFBII hydrophobin: nature of the adhesion energy, Langmuir 27 (2011) 4481–4488. [9] J.G.H. Wessels, Hydrophobins: proteins that change the nature of the fungal surface, in: R.K. Poole (Ed.), Advances in Microbial Physiology, vol. 38, Academic Press, 1996, pp. 1–45. [10] H.B. Wösten, J.H. Wessels, Hydrophobins, from molecular structure to multiple functions in fungal development, Mycoscience 38 (1997) 363–374. [11] H.A.B. Wosten, Hydrophobins: multipurpose proteins, Annu. Rev. Microbiol. 55 (2001) 625–646. [12] J. Hakanpaa, M. Linder, A. Popov, A. Schmidt, J. Rouvinen, Hydrophobin HFBII in detail: ultrahigh-resolution structure at 0.75 A acta crystallographica. section d, Biol. Crystallogr. 62 (2006) 356–367. [13] A.H. Kwan, I. Macindoe, P.V. Vukaˇsin, V.K. Morris, I. Kass, R. Gupte, A.E. Mark, M.D. Templeton, J.P. Mackay, M. Sunde, The Cys3–Cys4 loop of the hydrophobin EAS is not required for rodlet formation and surface activity, J. Mol. Biol. 382 (2008) 708–720. [14] M.J. Kershaw, N.J. Talbot, Hydrophobins and repellents: proteins with fundamental roles in fungal morphogenesis, Fungal Genet. Biol. 23 (1998) 18–33. [15] M.B. Linder, G.R. Szilvay, T. Nakari-Setala, M.E. Penttila, Hydrophobins: the protein-amphiphiles of filamentous fungi, FEMS Microbiol. Rev. 29 (2005) 877–896. [16] H.B. Wösten, K. Scholtmeijer, Applications of hydrophobins: current state and perspectives, Appl. Microbiol. Biotechnol. 99 (2015) 1587–1597. [17] M.B. Linder, Hydrophobins: proteins that self assemble at interfaces, Curr. Opin. Colloid Interface Sci. 14 (2009) 356–363. [18] M.S. Grunér, G.R. Szilvay, M. Berglin, M. Lienemann, P. Laaksonen, M.B. Linder, Self-assembly of class II hydrophobins on polar surfaces, Langmuir 28 (2012) 4293–4300. [19] J.G.H. Wessels, Developmental regulation of fungal cell wall formation, Annu. Rev. Phytopathol. 32 (1994) 413–437. [20] S. Askolin, M. Linder, K. Scholtmeijer, M. Tenkanen, M. Penttila, M.L. de Vocht, H.A.B. Wosten, Interaction and comparison of a class I hydrophobin from Schizophyllum commune and class II hydrophobins from Trichoderma reesei, Biomacromolecules 7 (2006) 1295–1301. [21] O.M.H. d. Vries, S. Moore, C. Arntz, J.G.H. Wessels, P. Tudzynski, Identification and characterization of a tri-partite hydrophobin from Claviceps fusiformis. A novel type of class II hydrophobin, Eur. J. Biochem. 262 (1999) 377–385. [22] A. Gravagnuolo, S. Longobardi; A. Luchini, M.S., Appavou, L. De Stefano, E., Notomista, L., Paduano, P. Giardina, Class I hydrophobin Vmh2 adopts atypical mechanisms to self-assemble into functional amyloid fibrils, 17 (2016) 954–964. [23] V. Lo, Q. Ren, C. Pham, V. Morris, A. Kwan, M. Sunde, Fungal hydrophobin proteins produce self-assembling protein films with diverse structure and chemical stability, Nanomaterials 4 (2014) 827–843. [24] M. Sunde, A.H.Y. Kwan, M.D. Templeton, R.E. Beever, J.P. Mackay, Structural analysis of hydrophobins, Micron 39 (2008) 773–784. [25] J. Bayry, V. Aimanianda, J.I. Guijarro, M. Sunde, J.-P. Latgé, Hydrophobins—unique fungal proteins, PLoS Pathog. 8 (2012) e1002700. [26] M. Khalesi, K. Gebruers, G. Derdelinckx, Recent advances in fungal hydrophobin towards using in industry, Protein J. 34 (2015) 243–255. [27] S. Houmadi, F. Ciuchi, M.P. De Santo, L. De Stefano, I. Rea, P. Giardina, A. Armenante, E. Lacaze, M. Giocondo, Langmuir-Blodgett film of hydrophobin protein from Pleurotus ostreatus at the air-water interface, Langmuir 24 (2008) 12953–12957. [28] Z.F. Wang, Y.J. Huang, S. Li, H.J. Xu, M.B. Linder, M.Q. Qiao, Hydrophilic modification of polystyrene with hydrophobin for time-resolved immunofluorometric assay, Biosens. Bioelectron. 26 (2010) 1074–1079. [29] K. Scholtmeijer, J. Wessels, H. Wösten, Fungal hydrophobins in medical and technical applications, Appl. Microbiol. Biotechnol. 56 (2001) 1–8.
Please cite this article in press as: B. Niu, et al., Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin, Colloids Surf. B: Biointerfaces (2016), http://dx.doi.org/10.1016/j.colsurfb.2016.10.048
G Model COLSUB-8230; No. of Pages 8 8
ARTICLE IN PRESS B. Niu et al. / Colloids and Surfaces B: Biointerfaces xxx (2016) xxx–xxx
[30] W.R. Yang, Q. Ren, Y.N. Wu, V.K. Morris, A.A. Rey, F. Braet, A.H. Kwan, M. Sunde, Surface functionalization of carbon nanomaterials by self-assembling hydrophobin proteins, Biopolymers 99 (2013) 84–94. [31] F. Sbrana, D. Fanelli, M. Vassalli, L. Carresi, A. Scala, L. Pazzagli, G. Cappugi, B. Tiribilli, Progressive pearl necklace collapse mechanism for cerato-ulmin aggregation film, Eur. Biophys. J. Biophys. Lett. 39 (2010) 971–977. [32] A.H.Y. Kwan, R.D. Winefield, M. Sunde, J.M. Matthews, R.G. Haverkamp, M.D. Templeton, J.P. Mackay, Structural basis for rodlet assembly in fungal hydrophobins, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 3621–3626. [33] J.P. Mackay, J.M. Matthews, R.D. Winefield, L.G. Mackay, R.G. Haverkamp, M.D. Templeton, The hydrophobin EAS is largely unstructured in solution and functions by forming amyloid-Like structures, Structure 9 (2001) 83–91. [34] X. Wang, H.P. Permentier, R. Rink, J.A.W. Kruijtzer, R.M.J. Liskamp, H.A.B. Wösten, B. Poolman, G.T. Robillard, Probing the self-assembly and the accompanying structural changes of hydrophobin SC3 on a hydrophobic surface by mass spectrometry, Biophys. J. 87 (2004) 1919–1928. [35] H. Fan, X.Q. Wang, J. Zhu, G.T. Robillard, A.E. Mark, Molecular dynamics simulations of the hydrophobin SC3 at a hydrophobic/hydrophilic interface, Proteins 64 (2006) 863–873. [36] L. Yu, B.H. Zhang, G.R. Szilvay, R. Sun, J. Janis, Z.F. Wang, S.R. Feng, H.J. Xu, M.B. Linder, M.Q. Qiao, Protein HGFI from the edible mushroom Grifola frondosa is a novel 8 kDa class I hydrophobin that forms rodlets in compressed monolayers, Microbiology (UK) 154 (2008) 1677–1685. [37] S. Hou, X.X. Li, X.Y. Li, X.Z. Feng, R. Wang, C. Wang, L. Yu, M.Q. Qiao, Surface modification using a novel type I hydrophobin HGFI, Anal. Bioanal. Chem. 394 (2009) 783–789. [38] W.F. Li, Y.B. Gong, H.J. Xu, M.Q. Qiao, B.L. Niu, Identification properties of a recombinant class I hydrophobin rHGFI, Int. J. Biol. Macromol. 72 (2015) 658–663. [39] B.L. Niu, Y.B. Gong, X.H. Gao, H.J. Xu, M.Q. Qiao, W.F. Li, The functional role of Cys3-Cys4 loop in hydrophobin HGFI, Amino Acids 46 (2014) 2615–2625. [40] B.L. Niu, D.D. Wang, Y.Y. Yang, H.J. Xu, M.Q. Qiao, Heterologous expression and characterization of the hydrophobin HFBI in Pichia pastoris and evaluation of its contribution to the food industry, Amino Acids 43 (2012) 763–771. [41] O.-J. Burrowes, G. Diamond, T.-C. Lee, Recombinant expression of pleurocidin cDNA using the pichia pastoris expression system, J. Biomed. Biotechnol. 2005 (2005) 374–384.
[42] M. Qin, L.K. Wang, X.Z. Feng, Y.L. Yang, R. Wang, C. Wang, L. Yu, B. Shao, M.Q. Qiao, Bioactive surface modification of mica and poly(dimethylsiloxane) with hydrophobins for protein immobilization, Langmuir 23 (2007) 4465–4471. [43] R. Eisert, L. Felau, L.R. Brown, Methods for enhancing the accuracy and reproducibility of Congo red and thioflavin T assays, Anal. Biochem. 353 (2006) 144–146. [44] H. LeVine Iii, 18] Quantification of -sheet amyloid fibril structures with thioflavin T Methods in Enzymology, vol. 309, Academic Press, 1999, pp. 274–284. [45] H. Levine, Thioflavine T interaction with synthetic Alzheimer’s disease (-amyloid peptides: detection of amyloid aggregation in solution, Protein Sci. 2 (1993) 404–410. [46] R. Sabate, S.J. Saupe, Thioflavin T fluorescence anisotropy: an alternative technique for the study of amyloid aggregation, Biochem. Biophys. Res. Commun. 360 (2007) 135–138. [47] P. Butko, J.P. Buford, J.S. Goodwin, P.A. Stroud, C.L. McCormick, G.C. Cannon, Spectroscopic evidence for amyloid-like interfacial self-assembly of hydrophobin Sc3, Biochem. Biophys. Res. Commun. 280 (2001) 212–215. [48] M. Portaccio, A.M. Gravagnuolo, S. Longobardi, P. Giardina, I. Rea, L. De Stefano, M. Cammarota, M. Lepore, ATR FT-IR spectroscopy on Vmh2 hydrophobin self-assembled layers for Teflon membrane bio-functionalization, Appl. Surf. Sci. 351 (2015) 673–680. [49] W.E. Klunk, R.F. Jacob, R.P. Mason, 19] Quantifying amyloid by congo red spectral shift assay, in: W. Ronald (Ed.), Methods in Enzymology, vol. 309, Academic Press, 1999, pp. 285–305. [50] H.A. Wosten, M.L. de Vocht, Hydrophobins, the fungal coat unravelled, Biochim. Biophys. Acta 1469 (2000) 79–86. [51] M.L. de Vocht, K. Scholtmeijer, E.W. van der Vegte, O.M.H. de Vries, N. Sonveaux, H.A.B. Wösten, J.-M. Ruysschaert, G. Hadziioannou, J.G.H. Wessels, G.T. Robillard, Structural characterization of the hydrophobin SC3, as a monomer and after self-assembly at hydrophobic/hydrophilic interfaces, Biophys. J. 74 (1998) 2059–2068. [52] F. Zampieri, H.A.B. Wösten, K. Scholtmeijer, Creating surface properties using a palette of hydrophobins, Materials 3 (2010) 4607–4625. [53] I. Macindoe, A.H. Kwan, Q. Ren, V.K. Morris, W.R. Yang, J.P. Mackay, M. Sunde, Self-assembly of functional, amphipathic amyloid monolayers by the fungal hydrophobin EAS, Proc. Natl. Acad. Sci. U. S. A. 109 (2012) E804–E811.
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Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin Baolong Niu a,b , Bingzhang Li a,b , Huifang Wang a , Ruijie Guo a,b , Haijin Xu c , Mingqiang Qiao c,∗∗ , Wenfeng Li a,b,∗ a Key Laboratory of Interface Science and Engineering in Advanced Materials, Taiyuan University of Technology, Ministry of Education, Taiyuan, 030024, PR China b College of Materials Science and Engineering, Taiyuan University of Technology, Taiyuan, 030024, PR China c State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, Tianjin, 300071, PR China
a r t i c l e
i n f o
Article history: Received 24 May 2016 Received in revised form 19 October 2016 Accepted 20 October 2016 Available online xxx Keywords: Hydrophobin Self-assembly Cys3–Cys4 loop Rodlets Amphipathic
a b s t r a c t We used protein fusion technology to transplant the Cys3–Cys4 loop of HGFI (a class I hydrophobin from Grifola frondosa) into a nonamyloidogenic hydrophobin HFBI (a class II hydrophobin from Trichoderma reesei) and replace the corresponding amino acids between Cys3 and Cys4 in this protein to identify whether this loop renders it amyloidogenic. Water contact angle (WCA) and X-ray photoelectron spectroscopy (XPS) measurements demonstrated that the mutant protein HFBI-AR could form amphipathic membranes by self-assembling at the hydrophilic mica and hydrophobic polystyrene surfaces. This property enabled the mutant protein to alter the surface wettabilities of polystyrene and mica as well as to change the elemental composition of siliconized glass. Atomic force microscopy (AFM) measurements indicated that, unlike class I hydrophobins, no amyloid-like rodlets were observed on the mutant protein HFBI-AR coated mica surface. Moreover, the Cys3–Cys4 region could not catalyze the mutant protein HFBI-AR to drive intermolecular association and formation of a cross- rodlet structure to resist depolymerization in organic solvents when it selfassembled at water–air interfaces. These results demonstrate that the Cys3–Cys4 loop is not the major determinant that initiates HGFI to form rodlets or account for the unique properties of the proteins. © 2016 Elsevier B.V. All rights reserved.
1. Introduction Hydrophobins belong to a class of small surface active proteins that are uniquely secreted by filamentous fungi [1–5]. These proteins can self-assemble into stable amphipathic films at hydrophobic–hydrophilic interfaces and are responsible for a wide variety of functions in fungal physiology [6–11]. Aside from having a characteristic pattern of eight cysteine residues (including two pairs of adjacent cysteines) and four intramolecular disulfide bonds, hydrophobins share little amino acid sequence homology [12–14]. All of these proteins can self-assemble at hydrophilic/hydrophobic
∗ Corresponding author at: Key Laboratory of Interface Science and Engineering in Advanced Materials, College of Materials Science and Engineering, Taiyuan University of Technology, Taiyuan, 030024, PR China. ∗∗ Corresponding author at: State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, No. 94 Weijin Road, Tianjin 300071, PR China. E-mail addresses: [email protected] (M. Qiao), [email protected] (W. Li).
interfaces into amphipathic films and improve surface wettability [8,15–18]. Based on the stability of the formed amphipathic monolayers and spacing of the cysteine residues, hydrophobins are categorized into two classes, I and II [19–22]. The membranes formed by class I hydrophobins are extremely robust and consist of amyloid-like fibrillar structures, known as rodlets, which can only depolymerize in strong acids, such as trifluoroacetic acid (TFA) [23–28]. By contrast, the films self-assembled by class II hydrophobins are not fibrillar and can be easily dissociated into detergent and alcohol solutions [29–31]. Based on the amino acid sequence comparison of class I and II hydrophobins, the most remarkable difference in the structures between class I and II hydrophobins is the presence of a disordered loop, such as Cys3–Cys4 and Cys7–Cys8 loops (polypeptide segments from Cys3 to Cys4 and Cys7 to Cys8, respectively) [32,33]. In the primary structure, class I hydrophobins have a much larger Cys3–Cys4 loop and the polypeptide segment between this loop displays much greater sequence variation (from 4 to 44 residues). In particular, the length of the inter-cysteine regions is highly variable. By contrast, class II hydrophobins display substantially
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more sequence similarity to one another and the lengths of the polypeptide segments between cysteines 3 and 4 (the Cys3–Cys4 loop) are fully conserved [13,15,24]. Protease digestion and mass spectrometry as well as hydrogen/deuterium exchange studies of class I hydrophobin SC3 show that the Cys3–Cys4 loop has the propensity to form an ␣-helix structure when it initially adheres to hydrophilic/hydrophobic interfaces [34]. Moreover, the molecular dynamics simulations of SC3 show that the Cys3–Cys4 loop is a major driver when SC3 self-assembles into rodlet structures [34,35]. However, when the number of amino acid residues in the Cys3-Cys4 loop was reduced to 15 residues in class I EAS hydrophobin, rodlet formation was not impaired and the surface activity and properties of rodlets were indistinguishable from the full length protein [13]. HGFI is a class I hydrophobin that is secreted by the edible mushroom Grifola frondosa. In a previous study, we found that HGFI could form typical rodlet structures on the mica surface [36]. The membranes formed by HGFI were robust and could not depolymerize in a hot detergent and alcohol solution [37,38]. Moreover, circular dichroism (CD) shows that self-assembly of HGFI at the water–air interface could give rise to significant secondary structural changes [38], which indicates that the interfacial assembly yields highly ordered -sheets to promote rodlet formation. To study the role of the Cys3–Cys4 loop in class I hydrophobin, we previously obtained a mutant protein HGFI-AR by replacing the Cys3–Cys4 loop of HGFI with the loops that are present in typical class II hydrophobin HFBI using protein fusion technology. The results show that the rodlet structures of HGFI are impaired and the membranes formed by the mutant protein become less robust after the Cys3–Cys4 loop is replaced [39]. Therefore, we speculate that the Cys3–Cys4 loop may play by major determinant in the formation of the rodlet structure of HGFI. To further investigate the contribution of the Cys3–Cys4 loop to the rodlet structure assembly and function of HGFI, we use protein fusion technology to transplant the Cys3–Cys4 region of HGFI into a nonamyloidogenic hydrophobin HFBI to identify whether this loop can render it amyloidogenic and change its physicochemical properties. 2. Materials and methods 2.1. Strains, vectors and reagents Pichia pastoris (P. pastoris) GS115, Escherichia coli (E. coli) DH5␣ and plasmid pPIC9 were purchased from Invitrogen (Beijing, China). pET-28a-hgfI and pMD19-T-hfbI were obtained from Prof. M.Q. Qiao (State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Nankai University, China). All of the restriction enzymes, polymerases, and ligases were purchased from TaKaRa (Dalian, China). DNA purification kits, DNA and protein markers were purchased from TaKaRa (Dalian, China). All of the primers used in this study were synthesized by Takara. Other reagents and chemicals were purchased from Sigma or TaKaRa.
Table 1 Primers used in the current study. Name
Sequence 5 –3
P1 P2 P3 P4 P5 P6 5 AOX 3 AOX
CGCTCGAGAAAAGAAGCAACGGCAACGGCAATG AGTGGAGGTAGACTCACAGCACTGGGGGTT CTCGTCGGTCTCACCTGCAAAGTCCCCTCC CGGAATTCAAGCACCGACGGCGGTCTG AACCCCCAGTGCTGTGAGTCTACCTCCACT GGAGGGGACTTTGCAGGTGAGACCGACGAG GACTGGTTCCAATTGACAAGC GCAAATGGCATTCTGACATCC
The restriction site XhoI in P1 and EcoRI in P4 are in underline; the Kex2 protease cleavage site is in bold.
as primers to obtain gene fragment A3. Fourth, the A1, A2 and A3 genes were used together as templates and P1/P4 as forward and reverse primers to generate the required mutant gene hfbI-ar by PCR. Then, the obtained hfbI-ar coding sequence was subcloned into the pMD19-T vector to give pMD19-T-hfbI-ar. The correct gene was confirmed by restriction enzyme digestion and DNA sequencing and was then cloned into the pPIC9 expression vector using XhoI and EcoRI restriction enzymes to obtain a recombinant vector, pPIC9-hfbI-ar. In the recombinant pPIC9-hfbI-ar vector, the hfbI-ar gene was located downstream of the AOX1 promoter, which can be induced using methanol as a sole carbon source. To ensure secretion of HFBI-AR, the ␣-factor secretion signal was placed upstream of the expressing gene, and the final recombinant vector pPIC9-hfbI-ar was verified by the XhoI and EcoRI cleavage and gene sequence analysis using the primer pairs 5 AOX/3 AOX. The hfbI-ar gene was transformed, and mutant protein HFBIAR was expressed and purified as described in Niu et al. [40]. The plasmid pPIC9-hfbI-ar was linearized with StuI and transfected into P. pastoris GS115 (His− ) cells by electroporation using a Bio-Rad gene pulser apparatus (25 F, 200 , and 2.0 kV) according to the manufacturer s instructions. Transformants were plated on minimal dextrose (MD) medium to screen for the His+ clones. Fifty His+ clones were selected and subsequently patched on MD and minimal methanol (MM) plates to determine the His+ Mut+ phenotypes. Colony PCR and sequencing were performed using the 5 AOX1 and 3 AOX1 primers to confirm the integration of hfbI-ar into the P. pastoris genome. The HFBI-AR expression levels from fifteen fastgrowing strains (His+ Mut+ ) were analyzed by RP-HPLC after they were induced by methanol supplementation to a final concentration of 0.5% at 24 h intervals for 5 days. The clone that expressed the highest level of HFBI-AR was chosen for large-scale production. A two-step process was used to purify the HFBI-AR protein. The first step was ultrafiltration using a hollow fiber membrane module with a 4-kDa molecular weight cut off and then lyophilized. RP-HPLC was used to further purify HFBI-AR with a Vydac C4 reversed-phase column. Introduction of the correct mutation HFBIAR was identified by 16% Tricine-SDS-PAGE, Western blotting and mass spectroscopy. 2.3. Water contact angle measurements
2.2. Production of mutant protein HFBI-AR The gene hfbI-ar was obtained by PCR using a four-step procedure, and the primers used are shown in Table 1. First, the pMD19-T-hfbI vector was used as a template and P1/P2 as forward and reverse primers to obtain gene A1. The KEX2 protease cleavage site following the XhoI restriction site was inserted at the 5 -terminus of P1. Second, pMD19-T-hfbI was also used as the template and P3/P4 as forward and reverse primers to obtain gene A2. An EcoRI restriction site sequence was added to the 5 end of P4. Third, the vector pMD18-T-hgfI was used as a template and P5/P6
The cleaned polystyrene and mica were incubated overnight in a 0.02 mg/mL HFBI-AR solution at 20 ◦ C. Then, the excess solution on the modified substrates was blown off with nitrogen gas. Finally, the protein coated surfaces were gently washed using water and dried with nitrogen gas again. To identify the stability of the film formed by HFBI-AR, the modified polystyrene was also washed using a 60% (v/v) ethanol solution or 2% boiling SDS solution. The WCAs of the modified and unmodified surfaces were measured with a 5-L water droplet using an optical contact angle meter (DSA100, Krus GmbH, Hamburg, Germany) at room temperature.
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The average WCA values were obtained from three measurements at different locations. 2.4. X-ray photoelectron spectroscopy (XPS) measurements The elemental compositions of bare and HFBI-AR (100 g/mL)coated siliconized glasses were analyzed with an XPS (Kratos Axis Ultra DLD) spectrometer employing a monochromated Al–Ka X-ray radiation source (h = 1486.6 eV), hybrid (magnetic/electrostatic) optics and a multi-channel plate and delay line detector (DLD). All of the XPS spectra were recorded using an aperture slot of 300 × 700 m. The survey spectra were recorded with a pass energy of 160 eV and high resolution spectra with a pass energy of 40 eV. 2.5. Atomic force microscope (AFM) measurements A 10-L aliquot of a 0.02 mg/mL HFBI-AR solution was deposited on a freshly cleaved mica substrate, and water was allowed to evaporate overnight in a covered container. The surface morphology of the HFBI-AR membrane was obtained using a NanoScope IIIa Multimode atomic force microscope (Veeco Instruments, NY, USA) in tapping mode at ambient atmosphere. The probe used contained silicon nitride cantilevers with a nominal force constant of 50 N/m. The scan rates were approximately 1 Hz. The damping ratio (set-point amplitude/free amplitude) was typically approximately 0.7–0.8. Analysis of the images was performed using a Scanning Probe Image Processor (SPIP; Image Metrology, Lyngby, Denmark).
Fig. 1. (a) Silver-stained Tricine-SDS–PAGE of purified HFBI-AR expressed in P. pastoris. M, molecular weight standards. Lane 1, fermentation supernatant of P. pastoris transformed with the pPIC9 vector. Lane 2, the lyophilisate of HFBI-AR concentrated/buffer exchanged by ultrafiltration. Lane 3, HFBI-AR purified by RP-HPLC. (b) Western blot analysis of HFBI-AR using a polyclonal antibody against HFBI. Lane 1, fermentation supernatant of P. pastoris transformed with the pPIC9 vector. Lane 2, the purified HFBI-AR.
CR and protein were incubated at room temperature for 30 min and shaken for 5 min prior to spectral analysis. Both CR and class I hydrophobin rHGFI were used as controls. 3. Results
2.6. Circular dichroism dpectropolarimetry 3.1. Expression and purification of HFBI-AR The secondary structure changes of HFBI-AR were measured by CD after it was self-assembled at an air-water interface. HFBIAR samples at a concentration of 50 g/mL in MQW were used in this study. The CD spectra of the assembled hydrophobins were obtained after vigorously shaking on a vortex mixer for 5 min. The spectra were recorded over the wavelength range from 190 to 250 nm on a Jasco J-715 CD spectrometer (Japan) using a 1-mm quartz cuvette. The temperature was kept at 25 ◦ C, and the sample compartment was continuously flushed with nitrogen gas. The samples’ spectra were the result of averaging 5 scans obtained using 5-s averaging per point, a bandwidth of 1 nm and a stepwidth of 0.5 nm. The spectrum of the reference solution without protein was used to correct for the background signal. 2.7. Thioflavin T (ThT) staining Purified HFBI-AR was dissolved in MQW and mixed with a stock ThT solution prepared in MQW. The effective concentrations of HFBI-AR and ThT in the mixture were 40 g/mL and 5 M, respectively. The sample and control (5 M ThT aqueous solution) for fluorimetry were performed using a Cary Eclipse Fluorescence Spectrophotometer (Varian Optical Spectroscopy Instruments, Mulgrave, Australia) before and after vortexing for 5 min. An emission scan from 450 to 600 nm was performed with an excitation wavelength of 435 nm. 2.8. Congo red (CR) binding assays CR binding was monitored by measuring the absorption spectra from 300 to 700 nm using a BIO-TEK Microplate Reader model (BioTek U.S., Winooski, VT, USA). Phosphate buffer (pH 4.0) was used to blank the instrument. Phosphate buffer (50 mM, pH 4.0) was used to dissolve CR and HFBI-AR. In the sample, the final concentrations of the protein and CR were 100 g/mL and 10 g/mL CR, respectively. Mixtures of
Gene A1 (86 bp) and A2 (159 bp) fragments were amplified by PCR using the pMD19-T-hfbI vector as a template. Gene A3 (126 bp) fragments were amplified by PCR using the pET-28a-hgfI vector as a template. A 311-bp HFBI-AR coding sequence was obtained using the A1, A2 and A3 gene sequences together as the template, and it was cloned into the P. pastoris expression vector pPIC9 to construct pPIC9-hfbI-ar. The hfbI-ar gene was located downstream of the AOX1 promoter, which means that HFBI-AR production can be induced by using methanol as the sole carbon source [41]. To ensure that full-length HFBI-AR was secreted to the extracellular environment, we designed a Lys-Arg linker to add after the XhoI site, which was recognized and specifically cleaved by the yeast KEX2 protease. The constructed plasmid pPIC9-hfbI-ar was linearized with StuI and transformed into the GS115 (His− ) strains using a Gene Pulser (Bio-Rad) apparatus (25 F, 200 X, 2.0 kV), as described in the manufacturer’s instructions. Fast growth of the selected transformants (His+ Mut+ ) was continued, as described by Niu et al. [40]. After being selected on MD and MM media plates, fifteen fast-growing transformants (His+ Mut+ ) were identified by PCR and sequencing using the 5 AOX/3 AOX primers. The clone with the highest HFBI-AR production (from the identified strains) was selected after the supernatants of the fermentation medium were analyzed by RP-HPLC. Ultrafiltration was used for the first step to remove salt and small impurities as well as to concentrate the extract. RP-HPLC was used for further purification. Both the lyophilized HFBI-AR after ultrafiltration and RP-HPLC purification were analyzed by 16% Tricine-SDS-PAGE. A clear band of approximately 9 kDa (coinciding with the expected size of HFBI-AR) was found in the selected transformant, but was absent from the control strains (P. pastoris transformed with pPIC9 vector) (Fig. 1a). Purified HFBI-AR was also identified by Western blot and mass spectrometry. The results showed that HFBI-AR could specifically bind to the polyclonal anti-HFBI antibody (Fig. 1b), and the molecular weight of this protein was 9599.19 Da, which is consistent with
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Table 2 The WCAa on the bare or HFBI-AR modified Polystyrene and mica surfaces.
Polystyrene Rinsed with 60% ethanol Rinsed with 2% SDS Mica
Bare surface (deg)
HFBI-AR(deg)
85 ± 2
36 ± 3 82 ± 2 84 ± 3 16 ± 2
7±2
The WCA values were averaged from three measurements at different locations. a n = 3.
the calculated value (9599.39 Da). As a result, we speculated that the target protein HFBI-AR was successfully expressed in P. pastoris and correctly secreted to extracellular medium without extra N-terminal amino acids. 3.2. Membrane formation and surface activity of the HFBI-AR protein Hydrophobins have a remarkable biophysical property, wherein they can self-assemble into amphipathic films at hydrophobic–hydrophilic interfaces to reverse their wettability. To detect whether the mutant protein HFBI-AR could convert the surface from hydrophilic to hydrophobic and vice versa, surface properties were probed with WCA measurements before and after HFBI-AR modification. The values of the WCA are shown in Table 2. The WCA of the bare polystyrene surface was 85 ± 2◦ , and this value dramatically decreased to 36 ± 3◦ with HFBI-AR coating on its surface. By contrast, the WCA results of the mica surface modified by HFBI-AR showed only a slight increase from 7 ± 2◦ to 16 ± 2◦ . Remarkably, the above results demonstrate that HFBI-AR is a surface active protein that contains hydrophobic and hydrophilic patches. As a result, once this protein coats the hydrophobic polystyrene surface, the hydrophobic region can adsorb onto its surface in response to hydrophobic interactions, which exposes the hydrophilic region to the outside to renders the polystyrene surface hydrophilic. In turn, when HFBI-AR self-assembles on the mica surface, the hydrophilic region faces the hydrophilic surface, leaving the hydrophobic region exposed to the outside, which subsequently made the mica surface hydrophobic. The WCA value of the mica surface only slightly increased after HFBI-AR-modified treatment, which could be explained as follows. First, as the reviewer suggested, the hydrophobin HFBI-AR might only partially cover the support surfaces; Second, once the mica surface is coated, the hydrophilic part of the first layer faces the surface and the hydrophobic patch of the first layer turns outside; then, the hydrophobic group of the second layer binds to the hydrophobic part of the first layer; finally, the hydrophilic part points outside. To further study the stability of the film formed by HFBI-AR, the HFBI-AR- modified polystyrene surfaces were thoroughly washed with 60% ethanol or 2% hot SDS. The WCA values dramatically increased from 36 ± 3◦ to 82 ± 2◦ and 84 ± 3◦ , which are close to that of the bare polystyrene surface. The restoration of the WCA values suggested that, unlike class I hydrophobins, the membrane formed by HFBI-AR was not as robust as that easily dissociated in 60% ethanol or 2% hot SDS. To explore the self-assembling properties of HFBI-AR, we also used XPS and AFM to detect the membrane formed by this protein on different surfaces. The XPS survey spectra and relative elemental composition of the siliconized glass surface before and after HFBIAR modification are shown in Fig. 2 and Table 3, respectively. A high intensity of O 1s and Si 2p signals and low intensity of the C 1s signal were detected on siliconized glass before HFBI-AR modification, and almost no S 2p s and N1 signals were observed in the spectra of the bare siliconized glass. After the surface was coated with HFBIAR, the elemental composition of Si 2p dramatically decreased from
Fig. 2. XPS spectra of siliconized glass surfaces before and after HFBI-AR modification.
Table 3 The relative elemental compositions of the bare and HFBI-AR modified siliconized glass surfaces. Spectra sample
C1s (%)
N1s (%)
O1s (%)
Si 2p (%)
S 2p (%)
Siliconized glass Siliconized glass + HFBI-AR
8.96 59.96
0.90 14.55
55.34 22.42
22.50 2.23
0.00 0.82
22.50% to 2.23% and the C 1s and N 1s signals became predominant in the spectra. In addition, a weak intensity of S 2p was also detected. Obviously, there is no Si element in HFBI-AR, which is only presents in siliconized glass. The high-intensity of the Si 2p and O 1s peaks represents a characteristic XPS spectrum of the siliconized glass surface. However, N and S elements only belong to the HFBI-AR protein family. Therefore, the increase in the N 1s and S 2p signals and decrease in the Si 2p signal confirm that HFBI-AR can form a membrane to cover the siliconized glass surface to a greater extent. Similar changes in peak intensity also occurred in the XPS spectra before and after hydrophobin HFBI was coated on the hydrophilic mica surfaces [42], which suggested that hydrophobin could also form a membrane on the hydrophilic mica surface, leading to an intensity decrease of the Si 2s and Si 2p peaks and a signal increase of the N 1s peak. However, when we rinsed HFBI-AR-coated siliconized glass with 2% hot SDS, the elemental composition reverted back to bare siliconized glass (data not shown). The result is in accordance with the conclusion obtained from the WCA measurements, which indicates that the membrane formed by HFBI-AR is not so robust that we cannot resist washing with hot SDS and ethanol. As already mentioned, the class I (not class II) hydrophobins present with rodlet structures when their dilute solution dries on a solid surface. AFM was also used in this study to observe the topography image of HFBI-AR. As stated in the Materials and methods section, when the HFBI-AR solution droplet dried on a freshly mica surface (Fig. 3a), only a compact monolayer of protein was clearly present and there was no rodlet structure observed. This membrane is different from the results found in our previous research that showed that typical mosaic rodlet structures (approximately 100 nm in length) were formed when a recombinant class I hydrophobin HGFI (rHGFI) solution was dried on a freshly cleaved mica surface (Fig. 3b). However, its topography image in AFM is
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Fig. 3. AFM images of different proteins dried onto mica; the image size is 1 × 1 m and the height scale is 10 nm for (a) HFBI-AR and (b) rHGFI.
Fig. 4. CD spectra of the mutant protein HFBI-AR in water (thick line) and after vigorous shaking (dotted line). The CD spectra were the averages of 5 scans.
similar to the membrane formed by recombinant class II hydrophobin HFBI (rHFBI) [38]. 3.3. Secondary structure changes upon self-assembly of HFBI-AR and rHGFI CD spectroscopy was used to study the secondary structural changes of HFBI-AR before and after self-assembly at the air-water interface. The spectra of monomeric HFBI-AR in water showed a maximum ellipticity that was close to 190 nm and a minimum that was close to 205 nm (Fig. 4 thick line). After vigorous shaking, only a slight decrease in the intensity of the spectrum was observed, which was due to the accumulation of protein at the liquid-air interface (Fig. 4 dashed line). Contrary to HFBI-AR, we previously observed that when the class I hydrophobin rHGFI solution was vortexed for 5 min, the assemblages of rHGFI acquired a predominately -sheet structure, which caused the minimum ellipticity to shift from 205 to 215–217 nm [39]. We speculate that the lack of stacked -sheets in solution-assembled HFBI-AR is present in the interfacially assembled rHGFI points to different assembly pathways between the two proteins. It has been reported that the aromatic dye ThT could specifically bind to stacked -sheet structures in amyloid fibrils to greatly
increase its fluorescence yield [43–45]. For this reason, it has been widely used to monitor amyloid formation [46]. The rodlet structure of class I hydrophobins shares many similarities with amyloid fibers, and the rodlets from SC3, EAS and rHGFI have been shown to bind ThT [13,38,47,48]. As a result, to examine whether the mutant protein HFBI-AR had the ability to self-assemble into amyloid-like structures, such as class I hydrophobins, the fluorescence probe ThT was utilized as described in the Materials and methods section. The fluorescence intensity was observed at 485 nm in the presence of a ThT and HFBI-AR-ThT mixed solution before and after vigorous vortexing (Fig. 5a), which could drive the intermolecular association and induce the protein solution to form rodlets by maximizing the air-water interfaces. Before vortexing, the fluorescence intensity value of the ThT/HFBI-AR mixed sample was 2-fold higher than that of a pure ThT water solution, which suggests that like other hydrophobins, HFBI-AR might contain some stacked -sheets in the monomeric structure or that the protein in the solution was not completely monomeric before testing. However, an increased fluorescence intensity of the ThT/HFBI-AR solution was hardly observed after vortexing for 5 min or longer (data not shown), which indicated that no substantial -sheet structure level formed when HFBI-AR selfassembled at the air-water interface. This phenomenon is different from our previously observed results that showed that when the recombinant protein rHGFI/ThT mixed solution was vortexed for 5 min (Fig. 5b), a pronounced increase in the fluorescence intensity was detected [38,39]. Moreover, -sheet structures dramatically increased after rHGFI self-assembled at the water-air interface. As a result, we speculate that the assembly pathways for rHGFI and HFBI-AR at the air-water interface were different. As in ThT, CR is a commonly used dye for amyloid detection because it can also bind to ordered stacks of -sheets and exhibits a significant shift in its absorption spectrum [49]. Because both CR and hydrophobins are negatively charged at neutral pH, the assay was performed at pH 4.0 to avoid electrostatic repulsion and prevent binding. The spectra of protein/CR and CR alone are shown in Fig. 6. Due to the limited extent of -sheets in BSA, this protein was used as a negative control. Meanwhile, the class I hydrophobin rHGFI, which could form rodlets, was also used as a positive control. Fig. 6 shows that the spectrum shifts from 490 nm to 530 nm when the rHGFI/CR mixed solution was vigorously vortexed. Like the negative control, a structural change at the air-water interface could not be observed from HFBI-AR, which indicates that the assemblages of this protein formation are not rodlets or sufficiently unstable to cause a spectrum shift.
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4. Discussion
Fig. 5. Fluorescence emission spectra of thioflavin T in the presence of HFBI-AR or control rHGFI. (a) The sample contained ThT alone (dashed line) or was unvortexed (dotted line) or vortexed (solid line) HFBI-AR. (b) The sample contained ThT alone (dashed line) or was unvortexed (dotted line) or vortexed (solid line) rHGFI.
Fig. 6. The adsorption spectra of different Congo red solutions.
Hydrophobins are small surface active proteins. Aside from a characteristic pattern of eight cysteine residues, there is little amino acid sequence similarity between these proteins. One of the major differences between class I and II hydrophobins is that in the primary structure, class I hydrophobins have a larger variable Cys3–Cys4 loop [2,50], which is the least conserved region in the protein. However, the corresponding loop in class II hydrophobins is smaller and conserved (11 residues). The secondary structure prediction suggests that part of the Cys3–Cys4 loop in class I hydrophobin has a high tendency to form a helix (as an anchor) to strongly bind to hydrophobic surfaces [51]. This prediction was confirmed by Wang; when their research team used various mass spectrometry techniques to monitor the structural changes of SC3, they found that the Cys3–Cys4 loop had a high affinity for Teflon and accompanied the formation of the ␣-helical structure to resist dissolution, which indicates that this loop is the key region driving intermolecular association and rodlet structure formation when SC3 self-assembles at hydrophilic/hydrophobic interfaces [34]. When class II hydrophobins self-assembled on the hydrophobic surface, the function of the Cys3–Cys4 loop differed and the smaller part bound much less at the surface [52]. Kwan et al. proposed a possible model for EAS rodlet formation [25], which suggests that the Cys3–Cys4 and Cys7–Cys8 disordered loop might “add on” to the -barrels to form an additional  sheet by H-bonds at air-water interfaces. However, when they removed 17 amino acid residues from 25 of the Cys3–Cys4 loops in EAS, the core structure of the mutant EAS was not essentially impaired and rodlets were still generated, which suggested that the Cys3–Cys4 loop is not central to EAS rodlet formation and the folding structure of the EAS monomer [13,53]. We previously replaced the amino acids between Cys3 and Cys4 of the class I hydrophobin HGFI from Grifola frondosa with those between Cys3 and Cys4 of the class II hydrophobin HFBI from Trichoderma reesei. Because replacement of the Cys3–Cys4 loop could impair the rodlet formation of HGFI [39], we conclude that the Cys3–Cys4 loop in HGFI may play an important role in initiating rodlet formation. To investigate the requirement for this loop in more detail, we inserted the Cys3–Cys4 region of HGFI into HFBI (a typical nonamyloidogenic hydrophobin) to identify whether this loop can render it amyloidogenic and change its physicochemical properties. Remarkably, the physicochemical properties of the mutant protein HFBI-AR are distinguishable from the class I hydrophobin rHGFI and other reported class I hydrophobins. Transplantation the Cys3–Cys4 loop is unable to trigger intermolecular associations of HFBI-AR along with stacked -sheet generation and render rodlet formation of this protein, as judged by ThT binding and AFM. Furthermore, compared to rHGFI, the membrane formed by the mutant protein HFBI-AR could not resist 60% ethanol and 2% hot SDS solubilization. This finding is consistent with the study by Kwan et al. [13], who found that deletion of 15 residues from the Cys3–Cys4 loop in EAS does not affect the physicochemical properties of its rodlet formation or change its surface activity. The previous reports showed that despite the very low level of sequence conservation, class I and II hydrophobins shared very similar folds. The three-dimensional structure showed that both class I and II hydrophobins had almost identical -barrel folds that consisted of four antiparallel -strands. This unique fold led the hydrophobins to form an amphipathic core structure [32,52]. Although the barrel portion of the class I hydrophobin fold was very similar to that of class II hydrophobin, the remainder of the structures were different. In class I hydrophobins, two disordered loops formed by the amino acid residues between Cys3–Cys4 and Cys7–Cys8 were present, but they were absent in class II
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hydrophobins. Moreover, the results of the triple-resonance NMR test showed that a two stranded -sheet in class I hydrophobins (EAS) to the outside of the -barrel was linked by the last disulfide, but the crystal structures of the class II hydrophobins (HFBI and HFBII) as solved by X-ray crystallography showed that in class II hydrophobins, the last disulfide links to an ␣-helix [13,24,52]. According to the possible model for class I hydrophobin rodlet formation, the two disordered loops could add on to the -barrels to form an additional  sheet at air-water interfaces, leading to class I hydrophobin rodlet structure formation [32]. As a result, the interaction between the Cys3–Cys4 and Cys7–Cys8 loops at airwater interfaces might play a central role in class I hydrophobin rodlet formation. Although part of the residues from the Cys3-Cys4 loop in class I hydrophobin EAS was deleted, rodlet formation was not hampered [13]. Site-directed mutagenesis and peptide experiments showed that inserting the Cys7-Cys8 loop of EAS into a nonamyloidogenic class II hydrophobin could enable it to form rodlets, is central to class I hydrophobin rodlet formation [53]. From the above results, we speculate that the major determinant that distinguishes the surface activity and self-assembly properties of class I and II hydrophobins is the amphipathic core region instead of the Cys3–Cys4 loop. Because the Cys3–Cys4 loop is not involved in the formation of this core region of class I hydrophobins, this flexible loop may only participate in the interaction of HGFI monomers with the interface, but is not responsible for triggering polymeric structure formation. As a result, when HFBI obtains the Cys3–Cys4 loop from HGFI, the physicochemical properties and structure are essentially unchanged. This observation also indirectly demonstrates why class I hydrophobins, which have this type of a non-conserved region in both the length and amino acid composition, can also display the same fold as well as polymeric rodlet structural similarities in morphology and dimensions. 5. Conclusions To investigate the contribution of the Cys3–Cys4 loop to the rodlet structure assembly and function of class I hydrophobins, we used protein fusion technology and obtained a mutant protein HFBI-AR by replacing the amino acids between Cys3 and Cys4 of class II hydrophobin HFBI with those that were between Cys3 and Cys4 of class I hydrophobin HGFI. Like native hydrophobins, HFBIAR retained amphiphilic properties, which could lead it to form amphipathic membranes at different surfaces and change their wettabilities. However, compared to class I hydrophobins, HFBI-AR could not self-assemble into rodlets and there was an accompanying increase in the -sheet conformational change in the secondary structure, which led to formation of a film in response to this protein becoming less stable. As a result, we speculated that the Cys3–Cys4 loop in class I hydrophobin HGFI is not the major determinant that initiates HGFI to form rodlets and distinguishes the surface activity and self-assembly properties of HGFI and HFBI. Conflict of interest The authors declare that they have no conflict of interest. Acknowledgments This research was financially supported by the National Natural Science Foundation of China (31170066) and Natural Science Foundation of Shanxi Province (2014021020-3, 2015021080), Scientific and Technological Innovation Programs of Higher Education Institutions in Shanxi (2015130), and Research Project Supported by Shanxi Scholarship Council of China (2015-033).
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References [1] M.L. de Vocht, I. Reviakine, H.A.B. Wösten, A. Brisson, J.G.H. Wessels, G.T. Robillard, Structural and functional role of the disulfide bridges in the hydrophobin SC3, J. Biol. Chem. 275 (2000) 28428–28432. [2] J. Hakanpää, A. Paananen, S. Askolin, T. Nakari-Setälä, T. Parkkinen, M. Penttilä, M.B. Linder, J. Rouvinen, Atomic resolution structure of the HFBII hydrophobin, a self-assembling amphiphile, J. Biol. Chem. 279 (2004) 534–539. [3] J. Wessels, O. De Vries, S.A. Asgeirsdottir, F. Schuren, Hydrophobin genes involved in formation of aerial hyphae and fruit bodies in schizophyllum, Plant Cell 3 (1991) 793–799. [4] V.K. Morris, Q. Ren, I. Macindoe, A.H. Kwan, N. Byrne, M. Sunde, Recruitment of class I hydrophobins to the air: water interface initiates a multi-step process of functional amyloid formation, J. Biol. Chem. 286 (2011) 9. [5] J.M. Kallio, M.B. Linder, J. Rouvinen, Crystal structures of hydrophobin HFBII in the presence of detergent implicate the formation of fibrils and monolayer films, J. Biol. Chem. 282 (2007) 28733–28739. [6] R.E. Beever, G.P. Dempsey, Function of rodlets on the surface of fungal spores, Nature 272 (1978) 608–610. [7] S.O. Lumsdon, J. Green, B. Stieglitz, Adsorption of hydrophobin proteins at hydrophobic and hydrophilic interfaces, Colloid Surf. B-Biointerfaces 44 (2005) 172–178. [8] E.S. Basheva, P.A. Kralchevsky, K.D. Danov, S.D. Stoyanov, T.B.J. Blijdenstein, E.G. Pelan, A. Lips, Self-assembled bilayers from the protein HFBII hydrophobin: nature of the adhesion energy, Langmuir 27 (2011) 4481–4488. [9] J.G.H. Wessels, Hydrophobins: proteins that change the nature of the fungal surface, in: R.K. Poole (Ed.), Advances in Microbial Physiology, vol. 38, Academic Press, 1996, pp. 1–45. [10] H.B. Wösten, J.H. Wessels, Hydrophobins, from molecular structure to multiple functions in fungal development, Mycoscience 38 (1997) 363–374. [11] H.A.B. Wosten, Hydrophobins: multipurpose proteins, Annu. Rev. Microbiol. 55 (2001) 625–646. [12] J. Hakanpaa, M. Linder, A. Popov, A. Schmidt, J. Rouvinen, Hydrophobin HFBII in detail: ultrahigh-resolution structure at 0.75 A acta crystallographica. section d, Biol. Crystallogr. 62 (2006) 356–367. [13] A.H. Kwan, I. Macindoe, P.V. Vukaˇsin, V.K. Morris, I. Kass, R. Gupte, A.E. Mark, M.D. Templeton, J.P. Mackay, M. Sunde, The Cys3–Cys4 loop of the hydrophobin EAS is not required for rodlet formation and surface activity, J. Mol. Biol. 382 (2008) 708–720. [14] M.J. Kershaw, N.J. Talbot, Hydrophobins and repellents: proteins with fundamental roles in fungal morphogenesis, Fungal Genet. Biol. 23 (1998) 18–33. [15] M.B. Linder, G.R. Szilvay, T. Nakari-Setala, M.E. Penttila, Hydrophobins: the protein-amphiphiles of filamentous fungi, FEMS Microbiol. Rev. 29 (2005) 877–896. [16] H.B. Wösten, K. Scholtmeijer, Applications of hydrophobins: current state and perspectives, Appl. Microbiol. Biotechnol. 99 (2015) 1587–1597. [17] M.B. Linder, Hydrophobins: proteins that self assemble at interfaces, Curr. Opin. Colloid Interface Sci. 14 (2009) 356–363. [18] M.S. Grunér, G.R. Szilvay, M. Berglin, M. Lienemann, P. Laaksonen, M.B. Linder, Self-assembly of class II hydrophobins on polar surfaces, Langmuir 28 (2012) 4293–4300. [19] J.G.H. Wessels, Developmental regulation of fungal cell wall formation, Annu. Rev. Phytopathol. 32 (1994) 413–437. [20] S. Askolin, M. Linder, K. Scholtmeijer, M. Tenkanen, M. Penttila, M.L. de Vocht, H.A.B. Wosten, Interaction and comparison of a class I hydrophobin from Schizophyllum commune and class II hydrophobins from Trichoderma reesei, Biomacromolecules 7 (2006) 1295–1301. [21] O.M.H. d. Vries, S. Moore, C. Arntz, J.G.H. Wessels, P. Tudzynski, Identification and characterization of a tri-partite hydrophobin from Claviceps fusiformis. A novel type of class II hydrophobin, Eur. J. Biochem. 262 (1999) 377–385. [22] A. Gravagnuolo, S. Longobardi; A. Luchini, M.S., Appavou, L. De Stefano, E., Notomista, L., Paduano, P. Giardina, Class I hydrophobin Vmh2 adopts atypical mechanisms to self-assemble into functional amyloid fibrils, 17 (2016) 954–964. [23] V. Lo, Q. Ren, C. Pham, V. Morris, A. Kwan, M. Sunde, Fungal hydrophobin proteins produce self-assembling protein films with diverse structure and chemical stability, Nanomaterials 4 (2014) 827–843. [24] M. Sunde, A.H.Y. Kwan, M.D. Templeton, R.E. Beever, J.P. Mackay, Structural analysis of hydrophobins, Micron 39 (2008) 773–784. [25] J. Bayry, V. Aimanianda, J.I. Guijarro, M. Sunde, J.-P. Latgé, Hydrophobins—unique fungal proteins, PLoS Pathog. 8 (2012) e1002700. [26] M. Khalesi, K. Gebruers, G. Derdelinckx, Recent advances in fungal hydrophobin towards using in industry, Protein J. 34 (2015) 243–255. [27] S. Houmadi, F. Ciuchi, M.P. De Santo, L. De Stefano, I. Rea, P. Giardina, A. Armenante, E. Lacaze, M. Giocondo, Langmuir-Blodgett film of hydrophobin protein from Pleurotus ostreatus at the air-water interface, Langmuir 24 (2008) 12953–12957. [28] Z.F. Wang, Y.J. Huang, S. Li, H.J. Xu, M.B. Linder, M.Q. Qiao, Hydrophilic modification of polystyrene with hydrophobin for time-resolved immunofluorometric assay, Biosens. Bioelectron. 26 (2010) 1074–1079. [29] K. Scholtmeijer, J. Wessels, H. Wösten, Fungal hydrophobins in medical and technical applications, Appl. Microbiol. Biotechnol. 56 (2001) 1–8.
Please cite this article in press as: B. Niu, et al., Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin, Colloids Surf. B: Biointerfaces (2016), http://dx.doi.org/10.1016/j.colsurfb.2016.10.048
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[30] W.R. Yang, Q. Ren, Y.N. Wu, V.K. Morris, A.A. Rey, F. Braet, A.H. Kwan, M. Sunde, Surface functionalization of carbon nanomaterials by self-assembling hydrophobin proteins, Biopolymers 99 (2013) 84–94. [31] F. Sbrana, D. Fanelli, M. Vassalli, L. Carresi, A. Scala, L. Pazzagli, G. Cappugi, B. Tiribilli, Progressive pearl necklace collapse mechanism for cerato-ulmin aggregation film, Eur. Biophys. J. Biophys. Lett. 39 (2010) 971–977. [32] A.H.Y. Kwan, R.D. Winefield, M. Sunde, J.M. Matthews, R.G. Haverkamp, M.D. Templeton, J.P. Mackay, Structural basis for rodlet assembly in fungal hydrophobins, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 3621–3626. [33] J.P. Mackay, J.M. Matthews, R.D. Winefield, L.G. Mackay, R.G. Haverkamp, M.D. Templeton, The hydrophobin EAS is largely unstructured in solution and functions by forming amyloid-Like structures, Structure 9 (2001) 83–91. [34] X. Wang, H.P. Permentier, R. Rink, J.A.W. Kruijtzer, R.M.J. Liskamp, H.A.B. Wösten, B. Poolman, G.T. Robillard, Probing the self-assembly and the accompanying structural changes of hydrophobin SC3 on a hydrophobic surface by mass spectrometry, Biophys. J. 87 (2004) 1919–1928. [35] H. Fan, X.Q. Wang, J. Zhu, G.T. Robillard, A.E. Mark, Molecular dynamics simulations of the hydrophobin SC3 at a hydrophobic/hydrophilic interface, Proteins 64 (2006) 863–873. [36] L. Yu, B.H. Zhang, G.R. Szilvay, R. Sun, J. Janis, Z.F. Wang, S.R. Feng, H.J. Xu, M.B. Linder, M.Q. Qiao, Protein HGFI from the edible mushroom Grifola frondosa is a novel 8 kDa class I hydrophobin that forms rodlets in compressed monolayers, Microbiology (UK) 154 (2008) 1677–1685. [37] S. Hou, X.X. Li, X.Y. Li, X.Z. Feng, R. Wang, C. Wang, L. Yu, M.Q. Qiao, Surface modification using a novel type I hydrophobin HGFI, Anal. Bioanal. Chem. 394 (2009) 783–789. [38] W.F. Li, Y.B. Gong, H.J. Xu, M.Q. Qiao, B.L. Niu, Identification properties of a recombinant class I hydrophobin rHGFI, Int. J. Biol. Macromol. 72 (2015) 658–663. [39] B.L. Niu, Y.B. Gong, X.H. Gao, H.J. Xu, M.Q. Qiao, W.F. Li, The functional role of Cys3-Cys4 loop in hydrophobin HGFI, Amino Acids 46 (2014) 2615–2625. [40] B.L. Niu, D.D. Wang, Y.Y. Yang, H.J. Xu, M.Q. Qiao, Heterologous expression and characterization of the hydrophobin HFBI in Pichia pastoris and evaluation of its contribution to the food industry, Amino Acids 43 (2012) 763–771. [41] O.-J. Burrowes, G. Diamond, T.-C. Lee, Recombinant expression of pleurocidin cDNA using the pichia pastoris expression system, J. Biomed. Biotechnol. 2005 (2005) 374–384.
[42] M. Qin, L.K. Wang, X.Z. Feng, Y.L. Yang, R. Wang, C. Wang, L. Yu, B. Shao, M.Q. Qiao, Bioactive surface modification of mica and poly(dimethylsiloxane) with hydrophobins for protein immobilization, Langmuir 23 (2007) 4465–4471. [43] R. Eisert, L. Felau, L.R. Brown, Methods for enhancing the accuracy and reproducibility of Congo red and thioflavin T assays, Anal. Biochem. 353 (2006) 144–146. [44] H. LeVine Iii, 18] Quantification of -sheet amyloid fibril structures with thioflavin T Methods in Enzymology, vol. 309, Academic Press, 1999, pp. 274–284. [45] H. Levine, Thioflavine T interaction with synthetic Alzheimer’s disease (-amyloid peptides: detection of amyloid aggregation in solution, Protein Sci. 2 (1993) 404–410. [46] R. Sabate, S.J. Saupe, Thioflavin T fluorescence anisotropy: an alternative technique for the study of amyloid aggregation, Biochem. Biophys. Res. Commun. 360 (2007) 135–138. [47] P. Butko, J.P. Buford, J.S. Goodwin, P.A. Stroud, C.L. McCormick, G.C. Cannon, Spectroscopic evidence for amyloid-like interfacial self-assembly of hydrophobin Sc3, Biochem. Biophys. Res. Commun. 280 (2001) 212–215. [48] M. Portaccio, A.M. Gravagnuolo, S. Longobardi, P. Giardina, I. Rea, L. De Stefano, M. Cammarota, M. Lepore, ATR FT-IR spectroscopy on Vmh2 hydrophobin self-assembled layers for Teflon membrane bio-functionalization, Appl. Surf. Sci. 351 (2015) 673–680. [49] W.E. Klunk, R.F. Jacob, R.P. Mason, 19] Quantifying amyloid by congo red spectral shift assay, in: W. Ronald (Ed.), Methods in Enzymology, vol. 309, Academic Press, 1999, pp. 285–305. [50] H.A. Wosten, M.L. de Vocht, Hydrophobins, the fungal coat unravelled, Biochim. Biophys. Acta 1469 (2000) 79–86. [51] M.L. de Vocht, K. Scholtmeijer, E.W. van der Vegte, O.M.H. de Vries, N. Sonveaux, H.A.B. Wösten, J.-M. Ruysschaert, G. Hadziioannou, J.G.H. Wessels, G.T. Robillard, Structural characterization of the hydrophobin SC3, as a monomer and after self-assembly at hydrophobic/hydrophilic interfaces, Biophys. J. 74 (1998) 2059–2068. [52] F. Zampieri, H.A.B. Wösten, K. Scholtmeijer, Creating surface properties using a palette of hydrophobins, Materials 3 (2010) 4607–4625. [53] I. Macindoe, A.H. Kwan, Q. Ren, V.K. Morris, W.R. Yang, J.P. Mackay, M. Sunde, Self-assembly of functional, amphipathic amyloid monolayers by the fungal hydrophobin EAS, Proc. Natl. Acad. Sci. U. S. A. 109 (2012) E804–E811.
Please cite this article in press as: B. Niu, et al., Investigation of the relationship between the rodlet formation and Cys3–Cys4 loop of the HGFI hydrophobin, Colloids Surf. B: Biointerfaces (2016), http://dx.doi.org/10.1016/j.colsurfb.2016.10.048