Letter pubs.acs.org/JPCL
Using SANS with Contrast-Matched Lipid Bicontinuous Cubic Phases To Determine the Location of Encapsulated Peptides, Proteins, and Other Biomolecules Leonie van ‘t Hag,†,‡,§ Liliana de Campo,∥ Christopher J. Garvey,∥ George C. Feast,§ Anna E. Leung,⊥ Nageshwar Rao Yepuri,⊥ Robert Knott,∥ Tamar L. Greaves,# Nhiem Tran,# Sally L. Gras,†,‡,▽ Calum J. Drummond,*,§,# and Charlotte E. Conn*,# †
Department of Chemical and Biomolecular Engineering and ‡Bio21 Molecular Science and Biotechnology Institute, The University of Melbourne, Parkville, Victoria 3010, Australia § CSIRO Manufacturing, Clayton, Victoria 3168, Australia ∥ Australian Centre for Neutron Scattering and ⊥National Deuteration Facility, Australian Nuclear Science and Technology Organisation (ANSTO), Lucas Heights, New South Wales 2234, Australia # School of Science, College of Science, Engineering and Health, RMIT University, Melbourne, Victoria 3001, Australia ▽ The ARC Dairy Innovation Hub, The University of Melbourne, Parkville, Victoria 3010, Australia S Supporting Information *
ABSTRACT: An understanding of the location of peptides, proteins, and other biomolecules within the bicontinuous cubic phase is crucial for understanding and evolving biological and biomedical applications of these hybrid biomolecule−lipid materials, including during in meso crystallization and drug delivery. While theoretical modeling has indicated that proteins and additive lipids might phase separate locally and adopt a preferred location in the cubic phase, this has never been experimentally confirmed. We have demonstrated that perfectly contrast-matched cubic phases in D2O can be studied using small-angle neutron scattering by mixing fully deuterated and hydrogenated lipid at an appropriate ratio. The model transmembrane peptide WALP21 showed no preferential location in the membrane of the diamond cubic phase of phytanoyl monoethanolamide and was not incorporated in the gyroid cubic phase. While deuteration had a small effect on the phase behavior of the cubic phase forming lipids, the changes did not significantly affect our results.
L
optimal region of the bilayer to retain their active conformation.14 It has been suggested that transmembrane proteins in the diamond cubic QIID phase will concentrate at the flat points to minimize the elastic energy of deformation.12 In a similar manner, it has been suggested that certain lipid additives, such as phospholipids, may segregate to preferred locations within the cubic phase.15 While modeling studies have indeed indicated that peptides, proteins and additive lipids might microphase separate and adopt a preferred location in the cubic phase, this has never been confirmed experimentally. Additionally, how the location and diffusion of bioactive molecules within the cubic phase evolves during in meso crystallization and drug delivery applications is also largely unknown.4,5 Herein we have used small-angle neutron scattering (SANS) to study the encapsulation of the transmembrane model peptide WALP2116 (Figure 1C) within the cubic phase of
ipid-based bicontinuous cubic phases are a robust, versatile, and inexpensive matrix for encapsulation of biomolecules such as peptides and proteins (Figure 1A,B).1,2 Because of their amphiphilic nature they are particularly suitable for the encapsulation of membrane proteins within a lipid bilayer environment.3 Applications of lipidic cubic phases include use as drug-delivery vehicles,4 for in meso membrane protein crystallization,5 and as biosensors6 and biofuel cells7 upon the encapsulation of enzymes. The influence of incorporated transmembrane proteins and peptides on the nanostructure of the cubic phase has been extensively investigated using small-angle X-ray scattering (SAXS).2,8−10 However, to our knowledge, the exact location of transmembrane proteins and peptides within the bicontinuous cubic phase unit cell has never been experimentally determined. While the bicontinuous cubic phases have zero mean curvature at the bilayer midplane, their Gaussian curvature is continuously varying from zero at the flat points, to the most negative curvature saddle points, Figure 1A,B.12 Lateral diffusion of hydrophobic proteins along the bilayer has been demonstrated13 potentially allowing some proteins to select an © 2016 American Chemical Society
Received: May 29, 2016 Accepted: July 11, 2016 Published: July 14, 2016 2862
DOI: 10.1021/acs.jpclett.6b01173 J. Phys. Chem. Lett. 2016, 7, 2862−2866
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The Journal of Physical Chemistry Letters
water, respectively,17 was synthesized in a fully hydrogenated (H-PE, Figure 1D) and fully deuterated (D-PE, Figure 1E) form. SANS measurements (Figure 2A,C) showed clear Bragg peaks for the cubic phases of both D-PE and H-PE in D2O. To achieve contrast matching, we calculated an appropriate mixing ratio for D-PE (volume: vD‑PE) and H-PE (volume: vH‑PE) based on a combined scattering length density (SLDM‑PE) equal to that of D2O (6.37 × 10−6 Å2) using eq 1 and the values in Table 1. SLDH‑PE and SLDD‑PE are scattering lengths of H-PE Table 1. Molecular Formula, Molecular Weight, Density and Neutron Scattering Length Density (SLD) for H-PE and DPE in D2Oa Figure 1. (A) Diamond cubic QIID phase minimal surface and (B) gyroid cubic QIIG phase minimal surface with arrows indicating the Gaussian curvature gradient from the flat points to the most negatively curved saddle points. (C) Secondary structure prediction of WALP21 with side chains shown as sticks.11 Chemical structures of (D) H-PE: hydrogenated phytanoyl monoethanolamide and (E) D-PE: deuterated phytanoyl monoethanolamide.
a
phytanoyl monoethanolamide (PE) in D2O. Perfect contrastmatching was required between the scattering of the lipid membrane and the aqueous solution to study the scattering of the peptide encapsulated within the lipid-based material. The lipid PE, which forms the gyroid QIIG and diamond QIID cubic phases at room temperature, at 10−30% w/w and >30% w/w
lipid
molecular formula
molecular weight (g mol−1)
density (g cm−3)
SLD (× 10−6 Å2)
M-PE (% v/v)
H-PE D-PE
C22O2H45N C22O2D45N
355.6 400.9
0.94 1.06
−0.02 7.44
14.5 85.5
Using eqs 1 and (2) the mixing ratio for M-PE was calculated.
and D-PE, respectively, both of which can be calculated using eq 2. The molecular volumes (vm) of H-PE and D-PE were calculated to be the same at 0.628 nm3 (Supporting Information S1.1). The bound scattering length of each atom is indicated as bc.
Figure 2. (A,C) SANS data; the intensity is shown on an absolute scale. (B,D) Corresponding SAXS curves on a relative intensity scale. Results shown are from (A,B) the diamond cubic QIID phase in excess D2O and (C,D) the gyroid cubic QIIG phase at 13% w/w D2O, all at 25 °C. In panel D, the small differences in peak positions could be caused by slight variations in hydration: QIIG lattice parameters were calculated to be 88−92 Å. The difference in hydration between these QIIG phases was calculated to be ∼3% v/v using the method of Squires et al.18 (equation 4,18 l = 13 Å19). 2863
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Figure 3. DSC for (A) H-PE and D-PE, (B) H-PE and D-PE in D2O, and (C) D-PE in H2O and D2O. SAXS results of H-PE and D-PE, both in H2O and D2O, lattice parameters in angstroms are shown for (D) the QIID phase and (E) the HII phase. H-PE in D2O was fluid isotropic at 55 °C. Samples in panels B−E were in excess hydration.
SLDM‐PE =
vH‐PE*SLDH‐PE + vD‐PE*SLDD‐PE vH‐PE + vD‐PE
QIIG phase. For the diamond cubic QIID phase, this corresponds to 1.2 transmembrane domains for each of the four flat points. This was calculated as 1.7 mol % for WALP21 (Supporting Information 1). The second concentration was approximately four times lower: ∼0.4 mol %. For WALP21 in the diamond cubic QIID phase SANS Bragg reflections in the ratio √2:√3, typical of Pn3m crystallographic symmetry, were observed at 0.4 mol % and 1.7 mol % peptide in M-PE (Figure 2A). Because scattering from the lipid cubic phase itself has been perfectly contrast-matched, these peaks must arise from peptide scattering, suggesting that the peptides have indeed been incorporated within the lipid bilayer environment of the cubic phase. Given that there was only one peptide molecule present per cubic phase unit cell (at 0.4 mol %), the data suggest that the WALP21 peptide might be diffusing over the surface of the QIID phase. The scattering intensity increased with increasing WALP21 concentration due to the additional hydrogenated material present (form factor of the peptide molecules). Corresponding SAXS curves (Figure 2B) showed a slight decrease in QIID lattice parameters with 1.7 mol % WALP21, represented by a slight shift of the Bragg peaks to higher q values, in agreement with previous work.19 Scattering simulations have shown that the intensity ratio of the first two Bragg peaks in scattering patterns of bicontinuous cubic phases can provide information on preferential locations according to the Gaussian curvature of the membrane (L. de Campo, T. Varslot, T. Castle, R. Mittelbach, M. Moghaddam, and S.T. Hyde, unpublished data). If guest molecules, here peptides, in an otherwise contrast-matched system were concentrated at the flat points, then the scattering would be dominated by the second rather than the first peak. For the diamond cubic QIID phase, the second peak would disappear if the guest molecules were concentrated at the most negative
(1)
n
SLDH/D‐PE =
∑i = 1 bci vm
(2)
The Bragg peaks for contrast-matched PE (M-PE) disappeared at the calculated mixing ratio 85.5:14.5 (D-PE:HPE v/v), demonstrating that the cubic PE membranes could be perfectly contrast-matched to D2O (Figure 2A,C). While small differences are expected in the local SLD between the head and tail of the amphiphile (Supporting Information Table S2), no Bragg peaks were observed in the contrast-matched sample of M-PE in D2O. Note that slight differences in phase behavior that were observed to exist between the hydrogenated and deuterated forms of PE, and for mesophases in H2O and D2O, will be discussed later in the manuscript. These differences are typically very small and are not predicted to significantly influence the SANS results presented herein. To our knowledge this is the first reported example of a perfectly contrast-matched cubic phase in D2O, allowing us to isolate the scattering from any encapsulated (hydrogenated) biomolecule. SANS was therefore used to examine the scattering of the transmembrane peptide WALP21 encapsulated within the contrast-matched cubic phases of PE. Previous research by us has shown that the purely hydrophobic WALP21 peptide does not have much effect on the diamond cubic QIID phase lattice parameters of PE, presumably due to minimal hydrophobic mismatch between the cylindrically shaped peptide (∼31 Å long) and the lipid bilayer thickness (∼26 Å).19 The WALP21 peptide concentration was specifically chosen to place one transmembrane domain of the WALP21 molecule on each of the 16 flat points in a unit cell of the gyroid cubic 2864
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samples in H2O. We have previously reported that the excess water point of H-PE is at 30% w/w,17 suggesting that the diamond cubic QIID phase can take 29% v/v of H2O and only 27% v/v of D2O. Lower effective hydration in D2O compared with H2O was also reported for the cubic phase of monoolein.20 In conclusion, we have demonstrated that perfectly contrastmatched cubic phases in D2O can be obtained by mixing fully deuterated and hydrogenated lipid at an appropriate ratio. This finding can be applied to determine the location of peptides, proteins, and other biomolecules in the bicontinuous cubic phase using SANS. Herein no enrichment was observed for the model peptide WALP21 at the flat points or most negative curvature saddle points of the diamond cubic QIID phase of PE, in contrast with what was suggested in several modeling studies. In addition, this result opens up the possibility of studying the conformation (and hence function) of amphiphilic proteins and peptides within a lipid bilayer environment. While deuteration had a small effect on the phase behavior of the cubic phase forming lipid, these changes did not adversely affect our interpretation of the data, which provides new insights into these complex systems.
curvature saddle points. The intensity ratio of the Bragg peaks of WALP21 at both concentrations in the contrast-matched diamond cubic QIID phase of M-PE in D2O (Figure 2A) was analyzed by fitting the first (√2) and second (√3) reflections with Gaussians. The ratios of the peak areas of the √2 and √3 reflections observed for WALP21 in the diamond cubic QIID phase were similar to the ratio for the noncontrast-matched H/ D-PE samples in D2O (17 to 18% c.f. 20−25%). This indicates that there was no enrichment of the peptide at the flat points or the most negative curvature saddle points, in contrast with what has been suggested in several modeling studies.12,15 All peak ratios and fitting of the data are shown in the Supporting Information 3. For WALP21 with the gyroid cubic QIIG phase we did not observe any SANS Bragg reflections and the scattering pattern at both concentrations was similar to M-PE (Figure 2C). The SANS scattering intensity increased with increasing WALP21 concentration due to the additional hydrogenated material (peptide) present. Because the corresponding SAXS curves (Figure 2D) demonstrate that the basic cubic phase structure was predominantly unaltered for these samples, the absence of SANS peaks indicates that the peptide was not incorporated within the gyroid cubic QIIG phase. We suggest that the high viscosity of the gyroid cubic phase samples, combined with the extremely low hydration (13% w/w), has precluded peptide uptake in this case. The effect is exacerbated by the high lateral pressure profile for branched chain lipids such as PE, which may also cause slow equilibration of the samples.19 In this case the peptide may be present as microcrystals or clusters of peptide coexisting with, but not incorporated within, the gyroid cubic phase. The effect of deuteration on the phase behavior of PE and for mesophases in H2O and D2O was studied using differential scanning calorimetry (DSC) and synchrotron SAXS (Figure 3). The DSC and SAXS data for H-PE and D-PE indicate that their phase behavior and excess water points were not identical. Specifically, as determined by DSC, the melting transition temperature of D-PE (−10 °C) was significantly lower than the melting transition temperature of H-PE (−6 °C). The devitrification transition of D-PE where the lipid crystallizes occurred at significantly higher temperatures (∼−40 to −20 °C) than for H-PE (∼−50 °C to −30 °C) (Figure 3A). The DSC data indicate, for samples with excess hydration, that the transition temperature from the diamond cubic QIID phase to the inverse hexagonal HII phase was 2 °C lower for D-PE than for H-PE. Additionally, the transition from hexagonal HII phase to fluid isotropic phase was 2 °C higher for D-PE than for H-PE in the same solvent (Figure 3B). The DSC results for H-PE in H2O and the repeat measurements for the samples with excess hydration are shown in the Supporting Information in Figure S4. The different source of the phytanyl chains as used for the synthesis of D-PE and H-PE may also contribute to the differences observed because the chains can come as a mixture of isomers. The excess water points of H-PE and D-PE were lower in D2O than in H2O, as shown by DSC (Figure 3C) and SAXS (Figure 3D,E). The transition temperature from the diamond cubic QIID phase to the hexagonal HII phase in excess D2O was consistently 1 to 2 °C lower than in H2O (Figure 3C). SAXS analysis also showed that lattice parameters of the diamond cubic QIID phase were always ∼0.5 Å lower for the samples in excess D2O than in H2O (Figure 3D). This indicates that the mesophases in excess D2O appear less hydrated than the
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EXPERIMENTAL METHODS Synthesis PE. H-PE was synthesized according to established protocols.17 For D-PE, phytanic acid-d39 and ethanolamine-d4 were synthesized according to published procedures.21−23 Phytanic acid-d39 was converted to phytanoyl chloride-d39 and then reacted with ethanolamine-d4 to form D-PE. An extensive description of the synthesis of D-PE can be found in Supporting Information 5. Sample Preparation. Samples with excess hydration for DSC and SAXS analysis were mixed in Eppendorf tubes and left to equilibrate for ∼7 days. Samples for SANS measurements were mixed in two Hamilton syringes (Hamilton Company, Reno, NV) with a coupler (TTP Labtech, Hertfordshire, U.K.) to control the water content precisely, except for M-PE with 0.4 mol % WALP21, which was mixed in a glass vial and then left to equilibrate for 12 h. For the QIIG phase, samples were made at 13% w/w hydration with respect to PE. Dif ferential Scanning Calorimetry. DSC was performed using a Mettler Toledo DSC 3 (Mettler Toledo, Melbourne, Australia). Samples were run at 10 °C min−1 under N2 purging at 40 mL min−1 and collected and analyzed using the STARe software package from Mettler Toledo. Small-Angle X-ray Scattering. SAXS analysis was performed at ANSTO (Lucas Heights, NSW, Australia) and the Australian Synchrotron (Melbourne, VIC, Australia).24,25 See Supporting Information 6 for additional information. Small-Angle Neutron Scattering. SANS measurements were made on QUOKKA26 at the ANSTO OPAL reactor with an incident wavelength of 5.0 Å and a wavelength spread of ∼10%. The sample detector distances were 1.3, 2, and 3 m. The data were normalized and presented on an absolute intensity scale. The Bragg peaks of all SANS and SAXS results were fitted using MATLAB as described and shown in Supporting Information 3.
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpclett.6b01173. 2865
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(1) Calculations for peptide concentrations. (2) Scattering length densities H/D-PE head and tail. (3) MATLAB analysis SANS and SAXS peaks. (4) DSC data for H-PE and D-PE in excess H2O and D2O. (5) Synthesis and characterization of D-PE. (6) SAXS methods. (PDF)
AUTHOR INFORMATION
Corresponding Authors
*C.J.D.: E-mail:
[email protected]. *C.E.C.: E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS We thank Prof. Raffaele Mezzenga for fruitful discussions on the project. We thank Yesim Gozukara for help with DSC measurements. We acknowledge support of the Bragg Institute, Australian Nuclear Science and Technology Organisation in providing neutron research facilities and the Australian National Deuteration Facility (partly supported by the National Collaborative Research Infrastructure Strategy (NCRIS) − an initiative of the Australian Government) for providing the chemical deuteration facilities used in this work. We acknowledge use of the SAXS/WAXS beamline at the Australian Synchrotron and we thank Drs. Adrian M. Hawley, Nigel M. Kirby, and Stephen T. Mudie for their assistance. We thank AINSE, Ltd for providing financial assistance (Award − PGRA) to L.v.H. to enable work at the Bragg Institute. S.L.G. is supported by the ARC Dairy Innovation Hub IH 120100005. C.E.C. is the recipient of an ARC DECRA Fellowship (DE160101281).
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REFERENCES
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