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pH induced changes in the surface viscosity of unsaturated phospholipids monitored using active interfacial microrheology Saba Ghazvini, Ryan Alonso, Nabil Alhakamy, and Prajnaparamita Dhar Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b02803 • Publication Date (Web): 11 Oct 2017 Downloaded from http://pubs.acs.org on October 12, 2017

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pH induced changes in the surface viscosity of unsaturated phospholipids monitored using active interfacial microrheology

Saba Ghazvini 1, Ryan Alonso 2, Nabil Alhakamy 3,4, Prajnaparamita Dhar *1,2

1. 2. 3.

Bioengineering Graduate Program, University of Kansas, Lawrence, KS 66046

Department of Chemical and Petroleum Engineering, University of Kansas, Lawrence, KS 66046.

Department of Pharmaceutics and Industrial Pharmacy, Faculty of Pharmacy, King Abdulaziz University, Jeddah, KSA 4.

Department of Pharmaceutical Chemistry, University of Kansas, Lawrence, KS 66047

*To whom correspondence should be addressed. Prajnaparamita Dhar (Telephone: +785-864-4969; Fax: +785864-4967; Email: [email protected])

Key Words: surface viscosity, unsaturated phospholipid, zwitterionic head-group, anionic head-group, Langmuir monolayer, pH, active interfacial microrheology

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Abstract Lipid membranes, a major component of cells, are subjected to significant changes in the pH depending on their location in the cell: the outer leaflet of the cell membrane is exposed to a pH of 7.4 while lipid membranes that make up late endosomes and lysosomes are exposed to a pH as low as 4.4. The purpose of this study is to evaluate how changes in the environmental pH within cells alter the fluidity of phospholipid membranes. Specifically, we studied pH induced alterations in the surface arrangement of monounsaturated lipids with zwitterionic head-groups (phosphoethanolamine (PE) and phosphocholine (PC)) that are abundant in plasma membrane, as well as anionic lipids (phosphatidylserine (PS) and phosphatidylglycerol (PG)) that are abundant in inner membranes, using a combination of techniques including surface tension vs. area measurements, interfacial microrheology and fluorescence/atomic force microscopy. Using an active interfacial microrheology technique, we find that phospholipids with zwitterionic head-groups show a significant increase in the surface viscosity at the acidic pH. This increase in surface viscosity is found to also depend on the size of the lipid headgroup, with a smaller headgroup showing higher increase in viscosity. The observed pH induced increase in viscosity is also accompanied by an increase in the cohesion pressure between zwitterionic molecules at acidic pH, and a decrease in the average molecular area of the lipids, as measured by fitting the surface pressure isotherms to well-established equations of state. Since fluorescence images show no change in the phase of the lipids, we attribute this change in surface viscosity to pH induced reorientation of the P--N+ dipoles that forms part of the polar lipid head-group, resulting in increased lipidlipid interactions. Anionic PG head-groups do not demonstrate this pH induced change in viscosity, suggesting that the presence of a net negative charge on the head-group causes electrostatic repulsion between the head-groups. Our results also show that active interfacial microrheology is a sensitive technique to detect minute changes in the lipid head-group orientation induced by changes in the local membrane environment, even in unsaturated phospholipids where the surface viscosity is close to the experimental detection limit.

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Introduction Lipid membranes form an integral part of cells. In addition to serving as a barrier between the extracellular and intercellular environment, as well as between the intracellular organelles, lipid membranes control many essential cellular processes such as cell signaling and cellular transportation. Since phospholipids form the fundamental building blocks of these lipid membranes, molecular interactions between phospholipid molecules is expected to influence the majority of cellular functions, due to alterations in the structure, lateral organization and dynamics of the phospholipids. Lateral organization of phospholipid membranes is also expected to be influenced by the presence of molecules in the surrounding environment as well as changes in the intracellular and extracellular environmental pH. For example, while the solution pH in the extracellular environment is maintained at 7.4, the interior of the cell is maintained at a pH of 7.0. Moreover, vesicles in the endocyclic pathway undergo changes in pH from 7.0 in early endosomes to pHs as low as 4.5-5.0 in late endosomes and lysosomes 1, 2, 3, 4, 5. One particular effect of changes in the intracellular pH on phospholipid membranes that remains unexplored is the effect of changes in intracellular pH on membrane fluidity. Membrane fluidity plays a crucial part in several cellular processes, and is modulated by changes in membrane packing and orientation. Extensive studies using deuterium NMR on phosphatidylcholine (PC) head-groups have established the ability of the polar P--N+ dipole in the lipids head-group to reorient themselves in response to the presence of charges in their vicinity, thus serving as a powerful sensor of electrostatic interactions between lipid molecules (often referred to as the “electrometer effect”). However, a fundamental understanding of how reorientation of the phospholipid head-group effects the fluidity of phospholipid membranes is currently not available. The presence of chemically different phospholipid head-group moieties in different cellular compartments introduces added complexities when exploring the effect of head-group reorientation on membrane fluidity. While the outer leaflet of eukaryotic cell membranes contain only zwitterionic lipid head-groups, such as phosphatidylcholine (PC) and phosphoethanolamine (PE), the location of anionic phospholipids such as phosphotidylserine (PS) in the inner leaflet of the plasma membrane ensures passive transport through the cell membrane by maintaining an electrostatic gradient

3, 4

. Further,

late endosomes and lysosomes have a higher composition of negatively charged phospholipids such as Bis Monoacylglycero Phosphate (BMP), which is a structural isomer of phosphotidylglycerol (PG). It is also important to note that while studies have shown the contributions of size and charge of the ions in the environment on the reorientation of the P--N+ dipole, the organization of the carbonyl chain or the glycerol part of the head-group were not affected by the presence of ions at the interface 6, 7, 8, 9. These observations suggest that chemically different phospholipid head-group moieties present in different cellular compartments may in fact have significant contributions to modulating membrane fluidity. Therefore, in order to obtain a complete understanding of the effect of pH changes in the cellular environment on lipid head-group re-orientations, the role of lipid head-group chemistry also needs to be considered, making this a major focus of this paper. Measuring the fluidity or surface viscosity of phospholipid membranes is non-trivial. This is especially true for unsaturated phospholipids that are the most abundant type of phospholipids in the cell membranes. This difficulty stems ACS Paragon Plus Environment

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from the inability to realize a completely isolated two-dimensional surface that is decoupled from the bulk sub-phase, as first identified by Saffmann and Dulbruck for lipid membranes. Detailed analysis of probe motion in lipid-laden interfaces suggest that this decoupling, quantified by the Boussinesq number B, should be much greater than 1 (B>>1). Such analysis suggests that most current rheometers are capable of accurately measuring surface viscosities as low as 10-7 Ns/m. However, surface viscosity of unsaturated phospholipids, which mostly remain in the liquid expanded phase, remain inaccessible to macroscopic rheometers, making measurements of their surface viscosity especially challenging10. Recent development of passive and active microrheology techniques, incorporating micron and sub-micron sized probes increase the sensitivity of the measurements to viscosities as low as 10-9(N-s/m), making the studies described here possible11, 12, 13 14. In this work, a previously described active microrheology technique 15 that analyzes the induced motion of nickel nanorods placed in a lipid monolayer, is used to monitor surface viscosity of four different unsaturated phospholipids at pH of 7.4 and 4.4. These two pHs are chosen to represent the limits in the pH encountered by phospholipid membranes in all cells, from the plasma membranes to late endosomes. Our choice of lipid head-groups is representative of the varying composition of the cellular membranes. Phosphatidylcholine (PC) head-group represents the most abundant zwitterionic phospholipid in the cell membrane. Phosphoethanolamine PE, also a zwitterionic lipid typically present in the inner leaflet of plasma membrane, is used here to compare the effect of the size of the head-group in the head-group reorientation process. PS and PG are used as representative anionic lipids, and PG is also used to model the composition of late endosomes. To limit our attention only on lipid head-group reorganization, and rule out any discrepancies in measurements resulting from differences in lipid chain length, the same mixed lipid alkyl chains (1-palmitoyl-2-oleoyl) are chosen in all four samples (table 1). Monounsaturated phospholipids with mixed alkyl chains of 1-palmitoyl-2-oleoyl form one of the main lipid components of eukaryotic cell membranes, and are therefore biologically relevant. Materials and Methods 1-hexadecanoyl-2-(9Z-octadecenoyl)-sn-glycero-3 phosphocholine (sodium salt) (POPC), 1-hexadecanoyl-2-(9Zoctadecenoyl)-sn-glycero-3-phosphoethanolamine(POPE), 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-L-serine (sodium salt) (POPS), and 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt) (POPG) were purchased from Avanti Polar Lipids, Alabaster, AL, as organic mixtures in chloroform. Texas Red® 1,2-dihexadecanoyl-sn-glycero-3phosphoethanolamine, triethylammonium salt, (TXR-DHPE) was purchased in the dried form from Life Technologies (Invitrogen) and dissolved in HPLC grade chloroform. The lipids were stored at -20 ᵒC when not in use. Salts sodium hydrogen phosphate (Na2HPO4), sodium dihydrogen phosphate (NaH2PO4), sodium chloride (NaCl), and potassium chloride (KCl) and calcium chloride dihydrate (CaCl2. 2H2O) were purchased from Fisher Scientific. All organic solvents used in this work were also purchased from Fisher Scientific. Phosphate Buffered Saline (PBS) at pH 4.4 and 7.4 with ionic strength of 185.56 mM was used for all experiments and was prepared using stoichiometric amounts of sodium hydrogen phosphate (Na2HPO4), sodium dihydrogen phosphate (NaH2PO4), sodium chloride (NaCl), and potassium chloride (KCl) in water (resistivity 18.2 MΩ/cm) using a Millipore Gradient System (Billerica, MA). A custom designed Langmuir trough, containing specially designed holders for electromagnetic coils, the purpose of which is explained below, was mounted on a standard Langmuir trough frame available from KSV-NIMA, Biolin ACS Paragon Plus Environment

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Scientific15. Supplementary figure S1 shows an image of this set-up. Two delrin barriers were used to alter the total surface area, by using a stepper-motor that is part of the frame. This whole set-up was mounted on a custom-modified fluorescence microscope from Nikon (Eclipse LV100). Surface pressure vs. area isotherms of phospholipids were recorded during the compression expansion isotherm by using a filter-paper Wilhelmy plate coupled to the Langmuir trough (KSV-NIMA, Biolin Scientific). The rate of compression used for the compression-expansion studies was 125mm/min. To initiate the surface pressure vs. area isotherm measurements in different sub-phase pHs, a 1 mg/ml organic solution of phospholipids (POPC, POPE, POPS, and POPG) in chloroform with 1 wt% of added TXR-DHPE dye molecules was first added dropwise at several locations on the buffer-air interface. The chloroform was allowed to evaporate for 20 minutes before compression was started. The change in the surface pressure with change in the mean molecular area was recorded using KSV-NIMA’s interface unit and accompanying software and saved for later analysis. Surface viscosity of the monolayer films was obtained by a previously described active microrheology technique 15

. Briefly, four home-built electromagnetic coils, oriented perpendicular to each other and placed in the magnet holder of

the modified Langmuir trough, were used to apply an external magnetic field (3 G – 100 G), which was used to orient nickel rods of size, diameter 300 nm and length 10-30 µm, placed on the lipid laden interface. The nickel nanorods were synthesized by electrochemical deposition of nickel into alumina templates [8-10], washed thoroughly by several centrifuging steps, re-suspended in a isopropanol:water mixture and added dropwise to the phospholipid covered interface. The capillary forces retain the nanorods at the interface, allowing the monitoring of the reorientation of individual nanorods. After addition of the nanorods to the monolayer surface, the monolayer was compressed to surface pressures 20 mN/m, 25 mN/m, and 30 mN/m using the barriers of the Langmuir trough, and images of the nanorod reorientation were recorded and saved for further analysis. To visualize the motion of individual nanorods, an extra-long working distance lens and a motorized stage were used to reduce the disturbances to the interface. The circular chamber between the electromagnets helped with controlling the drift of the interface, while the channels allowed free flow of the lipids in and out of the contained area, as shown in the supplementary data (Figure S1A). The drift was further controlled by using a custom designed microscope cover with a built-in window to monitor the position of the objective with respect to the air-water interface. Our set-up also allowed us to switch between bright field and fluorescence mode, to simultaneously obtain fluorescence images of the monolayer films and obtain information regarding existence of molecules in different monolayer phases. A uniformly bright area in the fluorescence images indicates that the lipids exist in a lipid expanded region, while appearance of dark domains in a bright background is indicative of lipid molecules in a coexistence region. Completely dark areas with bright boundaries are indicative of lipid molecules in lipid condensed phases. Finally, in order to obtain further information on any differences in lateral organization of zwitterionic vs. anionic lipids induced by a decrease in pH, we also obtained atomic force microscopy images of POPC and POPG lipid monolayers transferred onto a mica surfaces for both pHs. For this, we used a custom built inverse Langmuir-Schaffer technique as described previously30. Briefly, a freshly cleaved mica substrate was placed on an aluminum holder with machined knifeedges made of Teflon. The whole set-up was initially cleaned and placed on the bottom of the Langmuir trough and kept submerged in the same environmental conditions as described for the previous experiments, throughout the compression cycle. The lipid monolayer was compressed to the desired pressure of 30 mN/m, after which the buffer was slowly aspirated ACS Paragon Plus Environment

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until the knife-edge cut the monolayer and allowed it to fall on the substrate. This whole process could be visualized, since the trough was mounted on our custom-designed microscope. This ensured correct transfer of the film. After the film was transferred, it was washed in salt-free buffer at the same pH to avoid any artifacts from drying of salt crystals and then allowed to dry under flow of compressed air, before imaging. All transferred monolayers were imaged at ambient temperature in air using a Veeco di Multimode V microscope. A J scanner with an X−Y scan range of 125 × 125 μm2 was used in tapping mode using antimony-doped silicon probes (Bruker Scientific) with a resonance frequency of 371 kHz. Images were collected at a scan rate of 1 μm/s at a resolution of 512 pixels/ line. The images were later flattened using the built-in software to compensate for sample tilt (any raised features were excluded from this flattening). Theoretical analysis Analysis of nanorod motion: The equations describing the reorientation of magnetic nanorods (length l, magnetic moment µom) under an externally applied magnetic field have been described in detail in previous publications

11, 15, 16, 17

. Briefly, the motion of

these magnetic rheological probes in a purely viscous monolayer may be described by Eqn. 4, 𝜇𝜇0 𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑚 = −𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙 3

𝑑𝑑𝑑𝑑 𝑑𝑑𝑑𝑑

(4)

where H is typically the externally applied magnetic field and 𝜑𝜑 the angle between the length of rod and the applied magnetic field direction. The assumption of a purely viscous system is valid for the unsaturated phospholipid films studied here, since

unsaturation in the alkyl chains prevent these phospholipid molecules from assembling into well-organized liquid condensed or solid phases. Espinosa et al. showed that POPC lipid monolayers held at 30 mN/m showed no elastic modulus and presented all characteristics of a two-dimensional purely Newtonian fluid31. The rod reorientation is experimentally measured and analyzed using the ImageJ tracking software, and the rod motion plotted as a function of time can be described by the following equation:

where the relaxation time,𝜏𝜏 =

𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙 3 𝜇𝜇0 𝑚𝑚𝑚𝑚

𝑡𝑡 tan(𝜑𝜑/2) = exp �− � 𝜏𝜏

(5)

, is obtained by fitting the tan (φ/2) values of rod reorientation data to equation 4 using

the origin 8.6 software. The magnetic moment of the rod can be written in terms of the magnetization, M, and the rod aspect ratio, l/r. Therefore, the relaxation time, 𝜏𝜏 can be written as: 𝜏𝜏 =

𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙 3 𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙 3 𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙 2 𝑓𝑓𝑟𝑟 𝜂𝜂𝑤𝑤 𝑙𝑙3 = = = ( ) 2 𝜇𝜇0 𝑚𝑚𝑚𝑚 𝜇𝜇0 𝑀𝑀𝑀𝑀𝑀𝑀 𝜇𝜇0 𝑀𝑀𝑀𝑀𝑟𝑟 𝑙𝑙𝑙𝑙 𝜇𝜇0 𝑀𝑀𝑀𝑀𝑀𝑀 𝑟𝑟

(6)

By substituting the average magnetization of the rod (11.9 C/s.m) obtained as a result of calibrations in water and glycerol 12

, the total drag coefficient experienced by the nanorod, was obtained from experimental measurements. Since the total ACS Paragon Plus Environment

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drag, fr, is a sum of both frictional drag from the bulk water, fw as well as the surface drag , fs, fr= fw + fs , fs can then be calculated by subtracting fw from the experimentally obtained fr. The subphase drag, fw, was taken to be equal to half that of the drag on a rod of radius r and length l (for l/r ≥ 20) rotating in a viscous fluid:

11, 12, 18

.

For each value of 𝜏𝜏, the relationship between the Boussinesq number, B, and fs for an incompressible interfacial film was

determined by using the analytical relationship described by Dhar et. al.. 12. These values of fs were then used to calculate the surface viscosity from the Boussinesq number, B. Calculation of the association parameters of the phospholipid films at the different pHs: Experimental surface pressure-area isotherms can be theoretically described using equations of state that give more information about the intermolecular interactions between the molecules at the air-water interface. In a series of publications, Volhardt and coworkers have derived the generalized form of the Volmer’s equation of state for a multicomponent insoluble Langmuir monolayer19, 20, 21.

Specifically, an equation of state was derived by Volhardt and

coworkers specifically to describe the fluid (liquid-expanded and gaseous) state of the insoluble molecules in the monolayer. This equation is based only on the equations for the chemical potential of the solvent in the bulk phase and in the surface layer. The detailed steps of the derivation are described by Fainerman and Volhardt. Here, we briefly describe the most relevant equations. The equation of state for a surface layer with any number of components with any geometry was derived as: 𝛱𝛱 = −

𝑘𝑘𝑘𝑘 (ln 𝑥𝑥0 𝜔𝜔0

+ ln 𝑓𝑓0 )

(7)

Where 𝑘𝑘 is Boltzmann constant and 𝜔𝜔0 is the molecular area per solvent molecule. Assuming the enthalpy contribution,

ln 𝑓𝑓0𝐻𝐻 is independent of molecular area, and corresponds to a liquid-expanded monolayer, cohesion pressure, 𝛱𝛱𝑐𝑐𝑐𝑐ℎ , describes the interaction between the components.

𝛱𝛱𝑐𝑐𝑐𝑐ℎ =

𝑘𝑘𝑘𝑘 ln 𝑓𝑓0𝐻𝐻 𝜔𝜔0

(8)

An increase in cohesion pressure indicates an increased density of packing at the interface due to strengthening intermolecular interactions between the head-groups of the amphiphilic molecules and an increased robustness of the monolayer 20, 22. Using a parameter θ = 𝜔𝜔/𝐴𝐴 , to represent the monolayer coverage, and allowing for surface associations

and dissociations of the molecules in the surface layer, Fainerman and Vollhardt have derived a thermodynamic equation to define the following equation of state for multicomponent Langmuir monolayers 19, 20. Π=

𝜔𝜔 𝐴𝐴 𝜔𝜔 𝜔𝜔0 (1−( )) 𝐴𝐴

𝑘𝑘𝑘𝑘( )

− Π𝑐𝑐𝑐𝑐ℎ

(9)

Where Π is the surface pressure, 𝜔𝜔0 is the molecular area per solvent molecule, 𝜔𝜔 is the average molecular area of insoluble species, k is the Boltzmann constant and T is the temperature

21, 23

. By fitting equation 9 to the experimentally obtained

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isotherms, the association degree, defined as 𝑛𝑛 =

𝜔𝜔 𝜔𝜔𝜊𝜊

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, and the cohesion pressure between phospholipid molecules, Π𝑐𝑐𝑐𝑐ℎ ,

can be obtained. Fainerman and Vollhardt showed that the values of the molecular areas of the molecules found from the fitting of experimental Π-A isotherms to this proposed model was found to be in agreement with independent grazing Xray diffraction experiments for various amphiphilic molecules. The unsaturated lipids studied here are in a liquid –expanded state, justifying our choice of using their proposed model for our equation of state. Further, since this equation does not involve any geometric parameters of the amphiphilic molecules, we used this equation of state to describe our experimental isotherms for the different lipids with differences in head group size. Results and Discussion Effect of change in pH on zwitterionic head-groups To monitor the effect of pH on lipid organization at the air-water interface, we initially monitored alterations in the interfacial properties of POPC, the most abundant unsaturated phospholipid in the cell. As a first step, the surface pressure, Π, of a POPC monolayer was measured as a function of the mean molecular area occupied by the lipid molecules,

and comparisons in the surface-pressure area isotherms were made for the two pHs of interest. Figure 1A shows that as the

area available to the POPC molecules is reduced, Π increased gradually from zero to a maximum collapse surface pressure of ~45 mN/m, regardless of the subphase pH. However, the shape of the curve (or compressibility modulus) shows that for

a more acidic subphase (pH 4.4), the surface pressure vs. area isotherm is shifted towards lower area per molecule, indicating a possible condensing effect at more acidic pH. It is also important to note that unlike saturated phospholipids, unsaturated phospholipids do not show any phase transitions, since they are expected to remain in the liquid expanded (LE) phase with no specified crystalline order 4. The existence of the lipid molecules in the LE phase throughout the compression cycle was confirmed by fluorescence imaging of the samples, acquired at both pH 4.4 and pH 7.4 (Supplementary Figure S2), where no appearance of dark liquid condensed (Lc) domains was seen through the compression process. Therefore, even though the surface pressure vs. area isotherms suggest a possible condensing of the lipid molecules, this effect is not enough to cause a change in the phase of the lipid molecules. The shift in the POPC isotherm was also explored further by carefully analyzing the surface pressure vs. area isotherm using theoretically described equation of state shown in Equation 9. The red lines in Figure 1A shows the theoretical fit to the experimental isotherm for both pHs. Panel B of figure 1 summarizes the corresponding parameters which were obtained as a result of fitting the experimental isotherm to the theoretical equation of state. Our results show the average molecular area of insoluble species (𝜔𝜔) decreased at pH 4.4 compared to pH 7.4, suggesting a higher packing density indicative of a condensing effect at more acidic pH. This idea was further supported by the observation that the cohesion pressure, Π𝑐𝑐𝑐𝑐ℎ , increased when the sub-phase pH was changed from 7.4 to 4.4. Increase in the cohesion pressure with decrease in sub-phase pH can be ascribed to a strengthening of the intermolecular interactions, possibly because of the

acidic pH induced condensing effect. We hypothesize that such a condensing effect would also cause an increase in the surface viscosity due to an increase in the packing density of the lipids.

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To verify our hypothesis, we measured the surface viscosity of the POPC monolayer for the two relevant pHs. Figure 2A shows the average value of the total drag force, while Figure 2B shows the resulting surface viscosity of the POPC monolayers at surface pressures of 20, 25, and 30 mN/m for both pH 7.4 and pH 4.4. These surface pressures were chosen to ensure that we are still away from collapse, but at high enough surface pressures that we would expect our instrument to be sensitive to the interfacial drag forces. Moreover, a surface of 30 mN/m is considered the bilayer equivalent pressure, and is representative of lipid packing at physiological conditions. Figure 2A shows that the total drag force was at least 3 times higher than the measured drag force experienced by the probe particles at a lipid-free control sample (0.14, indicated as a line on the graph) for the lipid monolayers at neutral pH, and at least an order of magnitude higher for the more acidic pH. These results suggest that while our setup was capable of differentiating contributions from the interfacial stresses vs. stresses from the bulk solution, the contributions from the interfacial stresses are most reliable for the lower pH, where the contributions from the interface are most significant. Figure 2B shows the surface viscosity values calculated from the interfacial drag as a function of the surface pressure and shows that at pH 7.4, the surface viscosity is independent of the surface pressure and is very close to the detection limit of our system (10-9 Nm/s). This observation is not surprising for POPC, an unsaturated lipid in the LE phase. In a previous publication using a saturated phospholipid with a very long LE phase, we have shown that the surface viscosity of phospholipids is independent of the surface pressure in the LE phase, and only shows surface pressure dependence after a phase transition to the liquid condensed (LC) phase or when the phospholipid molecules exist in a LE-LC coexistence state15. More interestingly, we find that unlike at pH 7.4, at pH 4.4, the surface viscosity increases with surface pressure, and shows a maximum increase of twenty times higher surface viscosity at the highest surface pressure studied (30 mN/m). This increase in surface viscosity further confirms that a decrease in pH causes a condensing of the lipid molecules at the air-water interface. High resolution atomic force microscopy images shows direct evidence of cluster formation in POPC films transferred onto a solid mica substrate (Figure 11 top panel). However, it is important to note that this clustering is not enough to cause a phase transition to LC phase. Fluorescence images of POPC at both pHs show no appearance of domains at the more acidic pH. Further, as expected for a phospholipid in the LE state, the values of surface viscosity was found to be orders of magnitude lower than a disaturated phospholipid of the same head-group and same tail length (DPPC) at the same surface pressure, indicating that this change in surface pressure is not due to a phase transition to a more crystalline order at the interface. In fact, previous studies have shown that viscosity of trans- or cis-unsaturated PC could not be detected (due to very low viscosity values) while saturated PC at the interface exhibited very high viscosity in the LC phase 13. Based on our results, we conclude that a change in pH to a more acidic environment causes a condensing of the phospholipid molecules at the interface. We attribute this pH induced condensing effect to lipid head-group re-orientation at the air-water interface. Extensive NMR studies have shown that the P--N+ dipoles that make up the lipid head-group can act as a local charge sensor or “molecular electrometer”. In the presence of ions and other charged species, including proteins, these lipid dipoles can undergo a reorientation. The primary driving force for such conformational change of the lipid head-group is typically due to attraction/repulsion between the P--N+ dipoles and the partial charges in the surrounding micro-environment

6, 7, 8, 9, 24

. For example, it has been shown that while the P--N+ dipole is approximately parallel to the

membrane surface or the air-water interface for PC head-groups, the addition or binding of cationic molecule causes the N+ end of the dipole to tilt towards the water phase, changing the orientation of the phosphate segment to incorporate a more ACS Paragon Plus Environment

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densely packed arrangement of lipid tails, as shown in the schematic in Figure 3A. The angle and degree of reorientation of the head-group depend on the magnitude and type of the foreign charge (positive or negative),6, 7, 8, 9, 24. Wang, et. al., have more recently demonstrated a similar head-group reorganization induced by charged nanoparticles, resulting in local reconstruction of phospholipid bilayer. 25. They further showed that such a head-group reorganization resulted in a more gel-like or fluid-like membrane property, depending on the charge of the nanoparticles. We propose that a similar reorientation is possibly induced by the more acidic pH environment. At the more acidic pH, the presence of excess H+ ions can reorient the N+ end of the phosphate moiety such that the phospholipid tails pack more densely, similar to the observed effects in case of cationic molecules in the vicinity of zwitterionic PC lipids. A denser packing would in turn cause an increase in viscosity of the lipid molecules at the interface. Indeed, evidence of increase in packing density detected through a decrease in average molecular area of lipids (𝜔𝜔) based on Volmer’s equation, as well as the order of magnitude increase in surface viscosity confirm that interfacial microrheology might in fact be a unique tool to easily detect this electrometer effect that was previously measured using high resolution NMR techniques for phospholipid members. To further prove that this pH induced change in surface viscosity is not unique to the PC head-group alone, we applied the experimental techniques described above to POPE lipids. While both PE and PC lipid head-groups are zwitterionic, the size of PE head-group is smaller than PC head-groups, since only three H atoms are attached to the N atom as compared to the three methyl groups attached to the N atom in PC lipids. Therefore, we expect that environmental pH induced changes in head-group orientation will lead to increased changes in lipid packing in this case. Figure 4A shows a representative surface pressure-area isotherm of POPE monolayer at pH 4.4 and 7.4. As expected, the surface pressure increases with mean molecular area, until the POPE monolayer collapses at surface pressure ~47 mN/m. Further, apart from a slight shift in the isotherm, changes in the pH did not show any significant changes in the isotherms. As done in case of POPC, the surface pressure-area isotherms at both pHs, 7.4 and pH 4.4, were fitted using the Volmer’s generalized equation (Figure 4A). As shown in Figure 4B, similar to POPC, the average molecular area of insoluble species (𝜔𝜔) decreased at pH 4.4 compared to pH 7.4, suggesting higher packing density of the molecule at the more acidic pH. The ratio of molecular area occupied by lipid to water, n, increased at pH 4.4 compared to pH 7.4. Furthermore, Π𝑐𝑐𝑐𝑐ℎ increased at acidic conditions. Since n accounts for associations between the amphiphilic molecules, larger n and Π𝑐𝑐𝑐𝑐ℎ values for POPE compared to POPC suggest higher interaction between POPE molecules at pH 4.4.

Figure 5A shows the total friction factor as a function of surface pressure for POPE monolayers at sub-phase pH of

7.4 and 4.4. Again, one finds that the total drag does not increase with surface pressure for the neutral pH, but increases dramatically for the more acidic pH. Similarly, Figure 5B shows that the surface pressure shows a statistically significant change in surface viscosity (pPOPC>POPS>POPG) in total change in viscosity. The smaller size of the PE head-group causes the maximum change in surface pressure, while electrostatic repulsion between the negatively charged POPG lipids demonstrates the least change in viscosity. A previous study comparing the surface dilatational and shear rheology of saturated phospholipids DPPC and DMPE showed that at the same surface pressure DMPE had a higher surface viscosity compared to DPPC27. This difference was attributed to the smaller size of the PE head-group 27. Similarly, a previous study comparing the surface dilatational rheology of saturated lipids with similar head-groups (DPPC vs. DPPG) showed that in general DPPG demonstrated more fluid-like behavior compared to DPPC 28. In order to confirm that the pH induced change in surface viscosity cannot be related to a change in phase transition, we performed fluorescence microscopy imaging on the lipid monolayer. As noted in Supplementary figures S2, our results show no change in the phase state of the lipids due to changes in sub-phase pH for all four lipid group species. In order to obtain high resolution images of the evidence of pH induced lipid condensing in zwitterionic vs. anionic lipids, we obtained atomic force microscopy images of POPC and POPG lipids transferred onto mica substrates. Our results, shown in Supplementary Information Figure S4 demonstrate the appearance of nano silos in case of zwitterionic POPC lipids transferred from the interface onto a solid substrate at the two different sub-phase pHs. Such raised features are less in number in case of anionic POPG lipids. While these raised features do not confirm pH induced condensing of the lipid molecules and may well be lipid vesicles that collapse from the monolayer surface at high surface pressures, these features are present only at the more acidic pH, and are less in number in the anionic POPG lipids. Moreover, previous studies on lipid bilayers have shown such features in a POPC bilayer subjected to similar changes in environmental pH [5]. Therefore, we conclude that these raised features may indeed be a result of clustering of lipid molecules due to the more acidic pH in the membranes’ vicinity, which in turn cause an increase in the surface viscosity of these unsaturated lipids Conclusion In conclusion, we have presented how changes in the pH in the cellular environment can alter the packing of model phospholipid membranes at neutral and acidic pH. Both zwitterionic (POPC, and POPE) and anionic (POPS, and POPG) mono-unsaturated phospholipids which contain no two-dimensional crystalline order at the air-water interface at ambient ACS Paragon Plus Environment

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temperature were used for this study. Overall, it was found that there was a significant increase of surface viscosity at pH 4.4 for POPC, POPE, and POPS with increasing surface pressure, an effect that was absent in case of POPG. Since this dramatic increase in surface viscosity was not accompanied by any detectable changes in the phase of phospholipids, we attribute this pH induced change in surface viscosity to a reorientation of the P-_N+ dipole of phospholipid head-group in response to the presence of foreign charged species in the surrounding environment. Such lipid head-group reorientation is expected to result in increased associations between the molecules 6, 7, 8, 9, 24. The increased association between molecules at more acidic pHs was confirmed for the zwitterionic lipids using Volmer’s generalized equation of state that describes the surface pressure vs. area molecule isotherm. The presence of charges on the head-group moiety prevented such reorientation, as was the case for anionic POPG lipids. pH induced changes in surface viscosity has not been reported before for unsaturated phospholipids, possibly due to the limitations of accurate measurements in case of fluid monolayers. Our surface viscosity measurements show that the surface viscosity of mono-unsaturated phospholipid monolayers were very low compared to previously measured values for saturated lipid monolayers with the same head-group moieties for PC and PE head-groups at similar surface pressures and available area 15. Interfacial microrheology is extremely sensitive to minute changes in the lateral arrangement of lipid molecules, even when they cannot be detected by many other techniques. Moreover, measuring the surface viscosity of the more fluid unsaturated phospholipid is non-trivial, and interfacial microrheology may serve as an important technique to better understand lateral restructuring of phospholipids in physiological relevant changes in the local membrane environment. While this study focused only on changes in the pH, our group is interested in obtaining a thorough fundamental understanding of how subtle environmental changes around biological membranes due to changes in pH, ionic strength, or presence of charged species such as proteins or even nanoparticles can alter interfacial rheology and interfacial packing of lipid films. Further, while this study focused only on single component systems, biologically relevant lipid mixtures are more complicated and consist of sphingolipids and cholesterol that are expected to introduce viscoelectic behavior as well. Therefore, future studies will also focus on understanding how more complex multicomponent lipid mixtures respond to changes in the microenvironment resulting in changes to the viscoelectic properties of the monolayers. The studies presented here are a first step towards this effort. We also hope that these results will motivate some of our readers to consider addressing this important fundamental question using other complimentary techniques. Ultimately, one also needs to translate the results from lipid monolayer viscosities to more biologically relevant lipid bilayer systems.

REFERENCES

1. Mosesson, Y.; Mills, G. B.; Yarden, Y. Derailed endocytosis: an emerging feature of cancer. Nature Reviews Cancer 2008, 8 (11), 835-850. 2. Sorkin, A.; von Zastrow, M. Signal transduction and endocytosis: close encounters of many kinds. Nature reviews Molecular cell biology 2002, 3 (8), 600-614. 3. Van Meer, G.; Voelker, D. R.; Feigenson, G. W. Membrane lipids: where they are and how they behave. Nature reviews molecular cell biology 2008, 9 (2), 112-124. 4. Alberts, B.; Bray, D.; Lewis, J.; Raff, M.; Roberts, K.; Watson, J. D. Molecular biology of the cell, 1994. Garland, New York, 139-194. 5. Suresh, S.; Edwardson, J. M. Phase separation in lipid bilayers triggered by low pH. Biochemical and biophysical research communications 2010, 399 (4), 571-574. ACS Paragon Plus Environment

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6. Akutsu, H.; Seelig, J. Interaction of metal ions with phosphatidylcholine bilayer membranes. Biochemistry 1981, 20 (26), 7366-7373. 7. Bechinger, B.; Seelig, J. Interaction of electric dipoles with phospholipid head groups. A deuterium and phosphorus-31 NMR study of phloretin and phloretin analogs in phosphatidylcholine membranes. Biochemistry 1991, 30 (16), 3923-3929. 8. Doux, J. P.; Hall, B. A.; Killian, J. A. How Lipid Headgroups Sense the Membrane Environment: An Application of 14 N NMR. Biophysical journal 2012, 103 (6), 1245-1253. 9. Scherer, P. G.; Seelig, J. Electric charge effects on phospholipid headgroups. Phosphatidylcholine in mixtures with cationic and anionic amphiphiles. Biochemistry 1989, 28 (19), 7720-7728. 10. Hermans, E.; Vermant, J. Interfacial shear rheology of DPPC under physiologically relevant conditions. Soft Matter 2014, 10 (1), 175-186. 11. Dhar, P.; Cao, Y. Y.; Fischer, T. M.; Zasadzinski, J. A. Active Interfacial Shear Microrheology of Aging Protein Films. Phys Rev Lett 2010, 104 (1), -. 12. Dhar, P.; Fischer, T. M.; Wang, Y.; Mallouk, T.; Paxton, W.; Sen, A. Autonomously moving nanorods at a viscous interface. Nano letters 2006, 6 (1), 66-72. 13. Kim, K.; Choi, S. Q.; Zasadzinski, J. A.; Squires, T. M. Interfacial microrheology of DPPC monolayers at the air– water interface. Soft Matter 2011, 7 (17), 7782-7789. 14. Kim, K.; Choi, S. Q.; Zell, Z. A.; Squires, T. M.; Zasadzinski, J. A. Effect of cholesterol nanodomains on monolayer morphology and dynamics. Proceedings of the National Academy of Sciences 2013, 110 (33), E3054-E3060. 15. Ghazvini, S.; Ricke, B.; Zasadzinski, J. A.; Dhar, P. Monitoring phases and phase transitions in phosphatidylethanolamine monolayers using active interfacial microrheology. Soft matter 2015, 11 (17), 3313-3321. 16. Dhar, P.; Cao, Y. Y.; Kline, T.; Pal, P.; Swayne, C.; Fischer, T. M.; Miller, B.; Mallouk, T. E.; Sen, A.; Johansen, T. H. Autonomously moving local nanoprobes in heterogeneous magnetic fields. Journal of Physical Chemistry C 2007, 111 (9), 3607-3613. 17. Dhar, P.; Fischer, T. M.; Wang, Y.; Mallouk, T. E.; Paxton, W. F.; Sen, A. Autonomously moving nanorods at a viscous interface. Nano Letters 2006, 6, 66-72. 18. Dhar, P.; Cao, Y.; Kline, T.; Pal, P.; Swayne, C.; Fischer, T. M.; Miller, B.; Mallouk, T. E.; Sen, A.; Johansen, T. H. Autonomously moving local nanoprobes in heterogeneous magnetic fields. The Journal of Physical Chemistry C 2007, 111 (9), 3607-3613. 19. Fainerman, V.; Vollhardt, D. Equations of state for Langmuir monolayers with two-dimensional phase transitions. The Journal of Physical Chemistry B 1999, 103 (1), 145-150. 20. Fainerman, B.; Vollhardt, D. Equation of state for monolayers under consideration of the two-dimensional compressibility in the condensed state. The Journal of Physical Chemistry B 2003, 107 (14), 3098-3100. 21. Fainerman, V.; Vollhardt, D. Surface pressure isotherm for the fluid state of langmuir monolayers. The Journal of Physical Chemistry B 2006, 110 (21), 10436-10440. 22. Vollhardt, D.; Fainerman, V.; Siegel, S. Thermodynamic and textural characterization of DPPG phospholipid monolayers. The Journal of Physical Chemistry B 2000, 104 (17), 4115-4121. 23. Kuo, C.-C.; Kodama, A. T.; Boatwright, T.; Dennin, M. Particle size effects on collapse in monolayers. Langmuir 2012, 28 (39), 13976-13983. 24. Marassi, F. M.; Macdonald, P. M. Response of the phosphatidylcholine headgroup to membrane surface charge in ternary mixtures of neutral, cationic, and anionic lipids: a deuterium NMR study. Biochemistry 1992, 31 (41), 1003110036. 25. Wang, B.; Zhang, L.; Bae, S. C.; Granick, S. Nanoparticle-induced surface reconstruction of phospholipid membranes. Proceedings of the National Academy of Sciences 2008, 105 (47), 18171-18175. 26. Gurtovenko, A. A.; Lyulina, A. S. Electroporation of asymmetric phospholipid membranes. The Journal of Physical Chemistry B 2014, 118 (33), 9909-9918. 27. Krägel, J.; Kretzschmar, G.; Li, J.; Loglio, G.; Miller, R.; Möhwald, H. Surface rheology of monolayers. Thin Solid Films 1996, 284, 361-364. 28. Wüstneck, R.; Enders, P.; Wüstneck, N.; Pison, U.; Miller, R.; Vollhardt, D. Surface dilational behaviour of spread dipalmitoyl phosphatidyl glycerol monolayers. PhysChemComm 1999, 2 (11), 50-61.

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Table 1. Name, synonyms, classification, pKa, and structure of the POPC, POPE, POPS and POPG monolayers which are characterized in this study.

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Figure 1A. Surface pressure vs area isotherm of POPC monolayer at pH 7.4 (square) and pH 4.4 (circle) lines (presented as average, N=3). The red line represents the fitting curve based on generalized Volmer’s equation for insoluble monolayers to POPC isotherms at both pH 4.4 and 7.4. Panel 1B w = Average molecular area of amphiphilic species at the interface. w/ w0 = n (accounts for associations between the amphiphilic molecules). 𝜋𝜋𝑐𝑐𝑐𝑐ℎ = Cohesion pressure between molecules [19].

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Figure 2A and 2B represent Changes in total friction drag and surface viscosity as a function of pH (4.4 and 7.4) and surface pressure (20, 25, and 30 mN/m) of POPC. Results are presented as mean and ±1.5SE (n >= 3). The line in figure 2A represents the value of total friction of half immersed rod (0.14) in water as control. At SP 30mN/m POPC has a significant increase in surface viscosity compared to pH 7.4 (p= 3). The line in figure 5A represents the value of total friction of half immersed rod (0.14) in water as control. At SP 30mN/m and pH 4.4, POPE has a significant increase in surface viscosity compared to pH 7.4 (p= 3). The line in figure 7A represents the value of total friction of half immersed rod (0.14) in water as control. POPG do not have a significant viscosity change between pH4.4 and pH7.4 or with increase of surface pressure.

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Figure 8A. Surface pressure vs area isotherm of POPS monolayer at pH 7.4 (square) and pH 4.4 (circle) lines (presented as average, N=3). The red line represents the fitting curve based on generalized Volmer’s equation for insoluble monolayers to POPS isotherms at both pH 4.4 and 7.4. Panel 8B w = Average molecular area of amphiphilic species at the interface. w/ w0 = n (accounts for associations between the amphiphilic molecules). 𝜋𝜋𝑐𝑐𝑐𝑐ℎ = Cohesion pressure between molecules [19].

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Figure 9A and 9B represent Changes in total friction drag and surface viscosity as a function of pH (4.4 and 7.4) and surface pressure (20, 25, and 30 mN/m) of POPS. Results are presented as mean and ±1.5SE (n >= 3). The red line in figure 9A represents the value of total friction of half immersed rod (0.14) in water as control. At SP 30 mN/m the surface viscosity of POPS at pH 4.4 is three times more than pH 7.4 (p