Structural requirements for membrane binding of human guanylate-binding protein 1

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Article

Reference

Structural requirements for membrane binding of human guanylate-binding protein 1

SISTEMICH, Linda, et al.

Abstract

Human guanylate-binding protein 1 (hGBP1) is a key player in innate immunity and fights diverse intracellular microbial pathogens. Its antimicrobial functions depend on hGBP1's GTP binding- and hydrolysis-induced abilities to form large, structured polymers and to attach to lipid membranes. Crucial for both of these biochemical features is the nucleotide-controlled release of the C terminally located farnesyl moiety. Here, we address molecular details of the hGBP1 membrane binding mechanism by employing recombinant, fluorescently labeled hGBP1, and artificial membranes. We demonstrate the importance of the GTPase activity and the resulting structural rearrangement of the hGBP1 molecule, which we term the open state.

This open state is supported and stabilized by homodimer contacts involving the middle domain of the protein and is further stabilized by binding to the lipid bilayer surface. We show that on the surface of the lipid bilayer a hGBP1 monolayer is built in a pins in a pincushion-like arrangement with the farnesyl tail integrated in the membrane and the N-terminal GTPase domain facing outwards. We suggest that [...]

SISTEMICH, Linda, et al . Structural requirements for membrane binding of human guanylate-binding protein 1. The FEBS Journal , 2021, vol. 288, no. 13, p. 4098-4114

PMID : 33405388

DOI : 10.1111/febs.15703

Available at:

http://archive-ouverte.unige.ch/unige:149956

Disclaimer: layout of this document may differ from the published version.

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guanylate-binding protein 1

Linda Sistemich¹ , Lyubomir Dimitrov Stanchev^2,3, Miriam Kutsch^1,5 , Aurelien Roux⁴, Thomas Gunther Pomorski€ ^2,3and Christian Herrmann¹

1 Faculty of Chemistry and Biochemistry, Physical Chemistry I, Ruhr-University Bochum, Bochum, Germany 2 Faculty of Chemistry and Biochemistry, Molecular Biochemistry, Ruhr University Bochum, Bochum, Germany 3 Department of Plant and Environmental Sciences, University of Copenhagen, Frederiksberg C, Denmark 4 Biochemistry Department, University of Geneva, Geneva, Switzerland

5 Department of Molecular Genetics and Microbiology, Duke University Medical Center, Durham, NC, USA

Keywords

GBPs; large GTPase; membrane binding;

membrane tethering; protein-protein interaction

Correspondence

Christian Herrmann, Physical Chemistry I, Ruhr-University Bochum, 44780 Bochum, Germany.

E-mail: chr.herrmann@rub.de (Received 17 August 2020, revised 25 November 2020, accepted 29 December 2020)

doi:10.1111/febs.15703

Human guanylate-binding protein 1 (hGBP1) is a key player in innate immunity and fights diverse intracellular microbial pathogens. Its antimicrobial functions depend on hGBP1’s GTP binding- and hydrolysis-induced abilities to form large, structured polymers and to attach to lipid membranes. Crucial for both of these biochemical features is the nucleotide-controlled release of the C terminally located farnesyl moiety. Here, we address molecular details of the hGBP1 membrane binding mechanism by employing recombinant, fluorescently labeled hGBP1, and artificial membranes. We demonstrate the importance of the GTPase activity and the resulting structural rearrangement of the hGBP1 molecule, which we term the open state. This open state is supported and stabilized by homodimer contacts involving the middle domain of the protein and is further stabilized by binding to the lipid bilayer surface. We show that on the surface of the lipid bilayer a hGBP1 monolayer is built in a pins in a pincushion-like arrangement with the farnesyl tail integrated in the membrane and the N-terminal GTPase domain facing outwards. We suggest that similar intramolecular contacts between neighboring hGBP1 molecules are responsible for both polymer formation and monolayer formation on lipid membranes. Finally, we show that tethering of large unilamellar vesicles occurs after the vesicle surface is fully covered by the monolayer. Both hGBP1 polymer formation and hGBP1-induced vesicle tethering have implications for understanding the molecular mechanism of combating bacterial pathogens.

Databases

Structural data are available in RCSB Protein Data Bank under the accession numbers:6K1Z, 2D4H.

Introduction

Interferon-induced human guanylate-binding proteins (hGBPs) are key players in the innate immune response against bacterial, viral, and protozoan

pathogens [1–18] and show antiproliferative and anti- tumorigenic activity [19–22]. The family of hGBPs consists of seven members and belongs to the dynamin

Abbreviations

AlF_X, aluminum fluoride; DSP, dynamin superfamily protein; FRET, Forster resonance energy transfer; GDP, guanosine diphosphate; GED,€ GTPase effector domain; GMP, guanosine monophosphate; GppNHp, 5⁰-guanylyl imidodiphosphate; GTP, guanosine triphosphate; GTPcS, guanosine 5’-O-(gamma-thio)triphosphate; GUV, giant unilamellar vesicle; hGBP1, human guanylate-binding protein 1; hGBP1fn, farnesylated human guanylate-binding protein 1; LG, large GTPase domain; LUV, large unilamellar vesicle; MD, middle domain.

1 The FEBS Journal (2021)ª2021 The Authors.The FEBS Journalpublished by John Wiley & Sons Ltd on behalf of

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superfamily based on their domain architecture and biochemical properties [23,24]. Human GBP1 is hith- erto the best characterized hGBP paralog. Its multido- main architecture comprises an N-terminal globular large GTPase domain (LG domain) followed by a purely helical onset that can be subdivided in the middle domain (MD) and the GTPase effector domain (GED), and a CaaX box motif at the very C terminus (Fig.1). In its resting state, hGBP1 arranges in a closed, safety pin-like structure with its MD facing away from the LG domain and its GED folding back over the whole length of the protein[25]. The compact and tight structure of hGBP1 is mainly mediated by electrostatic interactions between the LG (R227, K228) and the GTPase effector domain (E556, E563, E568, E575) [25–28]. Upon binding of GTP, the LG domain undergoes conformational changes which allow dimerization of two hGBP1 molecules via their LG domains [26]. Formation of the LG domain interface results in intramolecular rearrangements, which disrupt the salt bridge network within each hGBP1 molecules and release the GEDs from the LG domains, ultimately allowing interactions between the GEDs of the hGBP1 molecules, which stabilize the dimer [27,29,30]. Dimer- ization stimulates hGBP1’s GTP hydrolysis, which is

exceptional among the dynamin superfamily proteins (DSPs) because after a first hydrolysis step the pro- duced GDP is hydrolyzed further to GMP [31–33].

This hGBP1-mediated GMP production was recently linked to inflammasome activation in the antibacterial host defense[34].

hGBP1 is unique among the seven hGBP paralogs by being farnesylated at its C-terminal CaaX box. In contrast, hGBP2 and hGBP5, which also harbor a CaaX motif at their C terminus, are geranylgerany- lated [35]. In cells, the prenylated hGBP paralogs are able to attach via their farnesyl or geranylgeranyl tail to different intracellular and bacterial membranes and guide nonprenylated hGBPs in a defined hierarchy to specific cellular compartments and bacterial pathogens [11,35–40]. Within this hierarchy, hGBP1 acts as a pioneer among all hGBPs. Previous studies have shown that binding of hGBP1 to host membranes, bacterial membranes, and artificial membranes is highly dependent on the presence and accessibility of the farnesyl tail [9,11,35,37,38,41,42]. In the nucleotide free, monomeric state the farnesyl moiety resides in a hGBP1- own hydrophobic pocket, strengthening the compact and closed structure of the protein [43] and shielding the farnesyl moiety from interactions with membranes.

The farnesyl moiety becomes accessible for membrane binding when released from the hydrophobic pocket, which is triggered following GTP binding. This nucleotide-controlled release was termed nucleotide-dependent farnesyl switch [38]. The farnesyl switch triggers not only interactions with membranes but also the formation of higher ordered hGBP1 structures [38,44].

Farnesylation of hGBP1 enables the protein to go beyond the state of dimerization during GTP hydrolysis, resulting in the formation of polymers consisting of up to 1000 hGBP1 molecules[44]. Polymerization is further stimulated by the interaction with bacterial lipopolysaccharide and is required for recognition and stable binding of gram-negative bacteria. Binding of hGBP1 to bacteria ultimately disrupts the integrity of the bacterial outer membrane, which recently explained several hGBP1-mediated antibacterial effects [42]. Interestingly, interaction of hGBP1 with host-like membranes occurs independently from polymerization and only requires the farnesyl switch. While in the nucleotide-free state hGBP1 is not capable to attach to host-like vesicles, binding of GTP induces the positioning of the farnesyl tail to an exposed conformation and allows interaction with the membrane [38].

Intriguingly, interactions of hGBP1 with host membranes seem not to interfere with the membrane integrity. Although hGBP1 is able to tether hGBP1-loaded vesicles in a GTP-dependent manner together, which LG domain

Middle domain α7-11 GTPase effector domain (GED)

α12-13

Farnesyl moiety

LG domain Middle domain α7-11

Farnesyl moiety GTPase effector domain (GED)

α12-13 A

B

Fig. 1.Domain architecture of the farnesylated human guanylate- binding protein 1 (hGBP1fn). (A) X-ray structure of the nucleotide unbound hGBP1_fn (PyMOL; Protein Data Bank ID code: 6K1Z).

hGBP1_fn consists of three domains: the large GTPase domain (LG domain; blue), the helical middle domain (MD;a7-11; yellow) and the helical GTPase effector domain (GED,a12-13; orange) with the farnesyl tail (black, circled) residing in its hydrophobic pocket (pink).

(B) Schematic representation of the X-ray structure with the different domains colored as in (A).

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notably also occurs in case of bacterial membranes, it fails to further modulate these host-like membranes [11,38]. This is in contrast to other DSPs that mediate host membrane fusion or fission after initial tethering [45–49].

Understanding the interaction of hGBP1 with host membranes further will therefore help to elucidate how hGBP1 distinguishes between host and pathogen membranes. Here, we explore further which structural requirements apart from the farnesyl switch are needed for hGBP1 to interact with host membranes.

Results

GTP hydrolysis enhances hGBP1 binding to membranes

We previously found that in the presence of GTP farnesylated hGBP1 (hGBP1fn) polymerizes and binds to artificial membranes. The assembly into polymers is strictly dependent on GTP hydrolysis and was only observed when offering the natural substrate GTP or GDP*AlF_X [38,44], a nucleotide complex known in GTPase research as a mimic for the GTP to GDP hydrolysis process. However, farnesylated hGBP1 failed to polymerize in the presence of the nonhy- drolyzable GTP-analogs GTPcS and GppNHp demonstrating that GTP binding alone is insufficient to induce polymerization [38]. In contrast, binding of hGBP1_fn to giant unilamellar vesicles (GUVs, used as artificial membrane system), occurred when offering GTP and GDP*AlFX, but also in the presence of GTPcS and GppNHp [38,39]. Here, we wanted to shed further light on the requirements for hGBP1fn’s membrane binding. Using the GUV binding assay, established in our previous study to observe binding of hGBP1_fn to single GUVs with confocal fluorescence microscopy, we noted that the fluorescence intensity of Alexa488-labeled hGBP1_fn colocalizing with rhodamine-labeled GUVs in the presence of GTP was higher compared to the intensity measured in the presence of GTPcS (Fig.2A). Since our previous studies demonstrated that hGBP1 undergoes complex conformational and structural changes regulated by the sequence of GTP binding, GTP hydrolysis and release of GTP hydrolysis products [26,30,33,44,50], we hypothesized that although GTP binding induces a conformation allowing hGBP1 binding to membranes, GTP hydrolysis might lead to a conformation of hGBP1 enhancing binding to membranes. In order to distinguish the effects that GTP binding and GTP hydrolysis have on hGBP1’s ability to bind membranes, we compared the intensities of hGBP1

colocalizing with GUVs, in the presence of GTPcS and GDP*AlFX, respectively. In comparison with the GTPcS-induced binding of hGBP1fn to GUVs, we observed a 50 times higher mean hGBP1fnfluorescence signal on the membrane when offering GDP*AlFX

(Fig.2B), demonstrating that upon GTP hydrolysis hGBP1 adapts a conformation which is more suitable for membrane binding.

LG domain-mediated dimerization is crucial for membrane attachment

The compact structure of hGBP1_fn in the nucleotide free state is mediated by electrostatic interactions between the LG and GTPase effector domain and supported by the farnesyl moiety residing in the hydrophobic pocket [27,29,43]. To release the tail and make it accessible for the surrounding, it is necessary that the protein undergoes intramolecular structural changes. Charge reversal in the LG domain (R227E/K228E, termed hGBP1-RK_fn) shifts the population of hGBP1 molecules from the closed conformation toward the open structure, facilitating dimerization, and polymerization [27,29,44]. To test whether disruption of the salt bridge network between the LG domain and the GED is sufficient to mediate binding of monomeric hGBP1 to membranes, we tested whether the mutant hGBP1-RK_fn is able to bind to membranes independently from GTP binding.

We found that in the monomeric, nucleotide-free, and GMP-bound states hGBP1-RKfn was not able to attach to GUVs (Fig.3A), suggesting that independent from a functional salt bridge network the farnesyl moiety remains in the hydrophobic pocket. In the presence of GTPcS, GTP and GDP*AlF_X, respectively, hGBP1-RK_fn attached to the membrane showing a similar behavior as the wild-type protein. We concluded that disruption of the salt bridge network is not sufficient to release the farnesyl tail for membrane binding but that at least binding of GTP is needed to induce the formation of LG:LG dimers, which are stabilized upon GTP hydrolysis and rearrangement of the GEDs. Notably, the RK mutant shows an increased GDPase activity compared to the wild-type protein (catalytic constants: hGBP1-RK: 0.28 min^-1; hGBP1:

0.00075 min^-1 [33]) reflecting, the formation of GDP- induced dimers. To test, if the GDP-induced dimer is also able to attach to membranes, the protein concentration was increased (10µ^M) with respect to the disso- ciation constant for the GDP-induced dimer (hGBP1- RK: K_D,GTPase: 0.74µ^M, K_D,GDPase: 1.93µ^M[51]) and observed that the RK mutant localizes at the

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membrane in the presence of GDP and GMP*AlF_X. Although the wild-type protein shows a lower affinity for the GDP-induced dimer[33], we could also observe binding to membranes in the presence of GDP and GMP*AlF_X (Fig.3B). In contrast to hGBP1-RK_fn, less vesicles were coated by wild-type hGBP1 (Fig.3C), reflecting the lower dimerization affinity and suggesting that in order to coat a vesicle, hGBP1 molecules stabilize each other.

To further challenge the importance of the LG domain-mediated dimerization interface for membrane binding, the mutant R244A was investigated which has compared to wild-type a weaker dimerization affinity [44,52]. R244 represents one of the two arginines that stabilize the formation of the LG domain interface. By replacing R244 by an alanine, the dimerization affinity of the nonfarnesylated mutant is decreased by a factor of 10 [52] and also the polymerization kinetics of the farnesylated protein are reduced by a factor of 20 [44]. We observed that weakening the dimer interface has consequences for GUV binding (Fig.3D,E). Although 2µ^M of

hGBP1-R244A_fn can be attracted to the membrane in the GDP*AlF_X bound state, binding of 2µ^M hGBP1-R244A_fn in the presence of GTP was not observed. By increasing the protein concentration 10 times to 20µ^M in the presence of GTP, the equilibrium was shifted toward the dimeric or oligomeric state. Under these conditions, hGBP1-R244A_fn was attracted to membranes in a GTP-dependent manner.

Thus, the protein-derived fluorescence on the membrane is increased from 0.01 a.u. for 2 µ^M to 24.5 a.u. for 20µ^M hGBP1-R244Afn (Fig.3D,E).

These results together demonstrated that in order to bind to host-like membranes dimerization of hGBP1 is required.

The GDP*AlF_X-induced protein coat on GUVs is stable toward detergent

Our observation that hGBP1 colocalized with only a part of the provided lipid vesicles in the presence of GDP and GMP*AlF_X (Fig.3B,C) not only reflected its low GDP-induced dimer affinity but also suggested

apo GTPγS GTP GDP*AlF_X

lipids rel. F(hGBP1)fn

A B

F(hGBP1)fn

Ordinate (μm)

hGBP1fn

0 10 20 30 0

100

40 200

0 10 20 30 0

100

40 200

0 10 20 30 0

100

40 200

0 10 20 30 0

100

40

200 apo

GTPγS GDP*AlF

x

0 100 200 2000

4000 ^****

****

Fig. 2.Nucleotide-dependent membrane binding of farnesylated hGBP1 to giant unilamellar vesicles (GUVs). (A) Binding of 2µ^MAlexa488- labeled hGBP1fn (green, upper panel) to rhodamine red labeled GUVs (magenta, lower panel) in the absence of nucleotide (apo) or in the presence of 500µ^M GTPcS 2 mMGTP or 250µ^MGDP*AlFX. Representative images from three independent experiments are shown. All scale bars equal 15µm. Images were gained under constant settings on the microscope. Below each image the hGBP1_fnfluorescence (F (hGBP1fn), [a.u.]) across the diameter of the GUV is shown. The spikes represent the protein-derived fluorescence intensity on the membrane[38]. Of note, in the absence of nucleotide hGBP1fnfailed to interact with the offered membranes, independent on the presence or absence of AlF_X(apo), as it was already shown in the latest studies[38,39]. (B) hGBP1_fnfluorescence (F(hGBP1_fn), [a.u.]) at the GUVs in the absence (apo) or presence of 500µ^MGTPcS or 250µ^MGDP*AlF_Xrelative to the background. Box plots show mean valuesSD for hGBP1fn fluorescence of 40–80 GUVs from three independent experiments for each nucleotide condition. Significance is determined with unpaired two tailed t-test in relation to experiment without nucleotide (apo). ****, P<0.0001.

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that in order to encapsulate a vesicle, membrane-attached hGBP1 molecules form a network to stabilize each other. To address the question whether the hGBP1_fn molecules are interacting among each other on the membrane apart from forming dimeric units, hGBP1_fn-loaded GUVs were dissolved with detergent.

The stability of the protein coat was challenged in the

presence of GDP*AlF_X, which induces the binding of hGBP1_fn more or less irreversibly to the membrane, and GTP, which creates a dynamic equilibrium of bound and unbound hGBP1_fn (Fig.4). To dissolve the vesicles after protein binding, 0.1 % Triton X-100 was added to the sample. In the case of GTP-induced binding the protein coat vanished as soon as the GUV was

apo GMP GDP GMP*AlF_X GTP GDP*AlF_X

A

GDP*AlFx

2 μM

GTP

2 μM 20 μM D

hGBP1-RKfn

hGBP1-R244A_fn

E

2 μ^M 20 μ^M 0

10 20 30 40 50

rel. F(hGBP1)fn

hGBP1-R244Afn

GDP GMP*AlF_X

hGBP1fn

B C

hGBP1

fn

hGBP1-RK

fn

hGBP1

fn

hGBP1-RK

fn

# GUV (%)

GDP GMP*AlFX

0 20 40 60 80 100

empty GUVs

protein-loaded GUVs GTPγS

Fig. 3.Nucleotide and concentration-dependent membrane binding of hGBP1-RK_fn, hGBP1_fnand hGBP1-R244A_fn to GUVs. A: Binding of 2µ^M(for apo, GMP, GTPcS, GTP, and GDP*AlFX) or 10µ^M(for GMP*AlFXand GDP) Alexa488-labeled hGBP1-RKfnto GUVs in the absence of nucleotide (apo) or in the presence of 250µ^MGMP, GMP*AlFX, GDP*AlFX, 2 mMGTP, GDP, or 500µ^MGTPcS. Representative images from three independent experiments are shown. All scale bars equal 15µm. B: Binding of 10µ^MAlexa488 labeled hGBP1_fnto GUVs in the presence of 2 mMGDP or 250µ^MGMP*AlFX. Representative images from three independent experiments are shown. All scale bars equal 15µm. C: Comparison of the number of protein-loaded (light gray) and empty (dark gray) GUVsSD in the presence of 10µ^MhGBP1fnor hGBP1-RK_fnand 2 mMGDP or 250µ^MGMP*AlF_X. For each condition, 40-50 GUVs from three independent measurements were evaluated.

D: Binding of 2µ^Mor 20µ^MAlexa488-labeled hGBP1-R244A_fnto GUVs in the presence of 250µ^MGDP*AlF_Xor 2 mMGTP. Representative images from three independent experiments are shown. All scale bars equal 15µm. For GTP-induced binding of different hGBP1-R244Afn

concentrations, microscopy settings were kept constant. E: hGBP1-R244A_fnfluorescence [a.u.] at the GUVs (relative to the background) of GTP-dependent binding of hGBP1-R244A_fnat the indicated concentrations in the presence of GTP. Box plots show mean valuesSD for relative fluorescence of 40 GUVs from three independent experiments for each concentration.

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disrupted (Fig.4A,B; Movie M1). In contrast, the protein coat formed by the GDP*AlF_X bound hGBP1_fn remained stable over time after dissolution of the vesicle. Although it was observable for different vesicles that the diameter of the coat was somewhat reduced after disrupting the vesicle, the protein coat did not collapse (Fig.4C,D; Movie M2), suggesting that hGBP1 molecules indeed stabilize each other once bound to the lipid vesicle.

The middle domain is required for stabilization of the dimer and exposure of the farnesyl tail For the polymer formation of farnesylated hGBP1, it was shown that the polymer reacts sensitively to small amounts of nonfarnesylated hGBP1. Nonfarnesylated hGBP1 cannot be incorporated in the polymer and abolishes polymerization by forming mixed hGBP1- hGBP1_fn-dimers [44]. Nevertheless, tissue culture studies demonstrated that the different hGBP paralogs interact with each other and that nonprenylated hGBPs can be recruited to intracellular membranes by

the prenylated paralogs with hGBP1 being the pioneer among all paralogs [35]. According to these findings, we wanted to test whether the farnesyl switch in the mixed dimer of farnesylated and nonfarnesylated hGBP1 is functional and if one farnesyl moiety is sufficient for the mixed dimer to bind to artificial membranes.

Therefore, hGBP1fn and hGBP1 were labeled with different fluorophores (hGBP1 with Alexa488 and hGBP1fn with Alexa647) and mixed in equimolar amounts. While nonfarnesylated hGBP1 alone did nei- ther bind to GUVs in the presence of GTP nor in the presence of GDP*AlF_X, binding of nonfarnesylated hGBP1 to GUVs was induced when farnesylated hGBP1 was added (Fig. 5A,C).

Next, we tested which interaction sites in the dimer have to be established to arrange hGBP1_fnin a conformation that is suitable for membrane binding. For this purpose, two truncated mutants were generated by further shortening hGBP1 from the C terminus. First, the GED (aa 482–592) was removed resulting in the LG- and middle domain (aa 1–481, termed hGBP1-DGED), 0 100 200 300 400 0.0

0.5 1.0 1.5

rel. F

time (s) 1.17 s 36.27 s 39.78 s 46.8 s 152.1 s 438.75 s

hGBP1fnlipids

C D

68.1 s 102.15 s 113.5 s 124.85 s 143.01 s 149.82 s

hGBP1fnlipids

A B

GTP

GDP*AlF_X 80 100 120 140 0.0

0.5 1.0 1.5 2.0

rel. F

time (s)

Fig. 4.GDP*AlF_X-induced networking of hGBP1_fnon the membrane. (A,C) 2µ^MAlexa488 labeled hGBP1_fn(green, upper panel) was bound to rhodamine red labeled GUVs (magenta, lower panel) in the presence of 2 mMGTP (A) or 250µ^MGDP*AlF_x(C). Afterward, Triton X-100 (0.1 %) was added to solubilize the vesicle membrane. Representative images from three independent experiments are shown. All scale bars equal 15µm. (B,D) Mean fluorescence intensities of hGBP1_fn(green) and the lipid channel (magenta) before and after solubilization of the membrane were recorded in the presence of 2 mMGTP (B) or 250µ^MGDP*AlF_x(D). Fluorescence intensities were normalized to the initial fluorescence (t=0 s). The timeline shows the duration of recording, and the arrow indicates the time point of membrane solubilization.

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and in a second truncation step, the complete helical part (middle domain and GED) was removed resulting in the isolated LG domain (aa 1–317, termed hGBP1- LG). Both truncated mutants on their own are not able to bind to membranes since they are lacking the C-terminal farnesyl moiety. hGBP1-DGED or hGBP1- LG (5µ^Meach) were mixed with hGBP1_fn (2µ^M) and membrane binding was induced with GDP*AlF_X (Fig.5B,C). While hGBP1-DGED colocalized with hGBP1_fn at the membrane (upper panel), hGBP1-LG was not recruited to the membrane, although hGBP1_fn was still binding (lower panel). To confirm that hGBP1fn interacts with all three mutants (hGBP1, hGBP1-DGED, hGBP1-LG) in the presence of GDP*AlFX, intermolecular FRET-measurements were performed showing mixed dimer formation for all the different constructs (Fig.5D). Based on these results, we concluded that the MD is required for the

formation of a hGBP1 dimer suitable for membrane binding.

hGBP1_fnforms a monolayer in upright orientation on the LUV surface

Apart from the simple membrane attachment a further characteristic of dynamin superfamily proteins (DSPs) is the ability to modulate membranes in terms of deformation, tubulation, fission, fusion, and tethering [24,45,53]. By now, for the farnesylated hGBP1 only tethering of artificial membranes could be demonstrated in a GTP-dependent manner [38]. In order to characterize hGBP1fn-induced vesicle tethering in a more quantitative way, we employed flow cytometry.

Different concentrations of hGBP1fn (0.5–15 µ^M) were incubated with a fixed concentration of LUVs (0.5 mgml^-1) in the presence of 250µ^M GDP*AlF_X,

Control hGBP 1

ΔGED LG

0 40 80

F(Alexa488)

lipids lipids C

hGBP1_fn trunc

GDP*AlFX

hGBP1_fn hGBP1

A

B

GTPGUV

D

ΔGEDLG

GUV 0 100 200

1.0 1.5 2.0

F/F0

Time (s) 300

Fig. 5.Colocalization of hGBP1fnwith different truncated mutants of hGBP1fnon GUVs. (A) Colocalization of 2µ^MAlexa488-labeled hGBP1 (marked with a green star) with 2µ^MAlexa647-labeled hGBP1_fn(marked with a cyan star) on rhodamine red labeled GUVs in the presence of 2 mM GTP or 250µ^MGDP*AlF_X. B: Colocalization of 5µ^M Alexa488-labeled hGBP1-DGED or 5µ^M Alexa488-labeled hGBP1-LG (both marked with a green star) with 2µ^MAlexa647-labeled hGBP1fn(marked with a cyan star) on rhodamine red-labeled GUVs in the presence of 250µ^MGDP*AlFX. All scale bars equal 15µm. For each condition (A,B), a representative image of three independent measurements is depicted to show binding qualitatively. (C) Protein fluorescence [a.u.] at the GUVs of Alexa488-labeled truncated mutants (hGBP1, hGBP1- DGED, hGBP1-LG) in the presence of 2µ^M Alexa647-labeled hGBP1_fn and 250µ^M GDP*AlF_X relative to the background. As control, the protein fluorescence at GUVs of 2µ^MAlexa488-labeled hGBP1 in the absence of hGBP1fn and in the presence of 250µ^M GDP*AlFXis depicted. Box plots show mean valuesSD of 40 GUVs from three independent measurements for each condition. (D) Representative fluorescence time courses (of three replicates) of intermolecular FRET of differently truncated hGBP1_fnmutants interacting with hGBP1_fn. Donor labeled truncated mutants (cyan: 2µ^MhGBP1, black: 5µ^MhGBP1-DGED, orange: 5µ^MhGBP1-LG; all marked with a green star) and 2µ^Macceptor-labeled hGBP1_fn(marked with a cyan star) were mixed in the presence of 250µ^MGDP*AlF_X. The change in fluorescence is relative to the initial fluorescence. The difference in the fluorescence increase is due to the different labeling efficiencies (see Material and Methods—Labeling of protein with fluorescent dye).

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and fluorescence intensities of membrane-bound hGBP1fn were measured after 1 min. In the presence of low protein concentrations (e.g., 0.5µ^M hGBP1fn), the LUV population showed a homogenous distribu- tion with a defined size (Fig.6A,B). With increasing protein concentration, the vesicle population changed by becoming more heterogenous, and a shift from small particles, like the single vesicles observed in the presence of low protein concentration, to larger particles was observed (Fig.6C,D). This change in the vesicle population was detected in the different fluorescence channels as well as in the forward and sideward scatter, which is independent from any fluorescence and a reliable measure of the vesicle size (Fig.6E). Based on the scatter plot in the presence of high hGBP1_fn concentration, two distinct regions were chosen to represent two different populations of vesicles (Fig.6E: orange and dark cyan). Utilizing fluorescent beads with a defined diameter (Fig.6F) a size estimation was assigned to the two different vesicle populations, showing that the vesicle population being present for all hGBP1_fn concentrations (Fig.6B,D:

orange) were approximately 200 nm in size and the second vesicle population showing up for high hGBP1_fn concentrations covers particles between 500 and 1000 nm in diameter (Fig.6D,E: dark cyan). We explained the shift in particle size in the presence of higher hGBP1_fn concentrations by the previously observed effect of liposome tethering induced by GTP binding [38]. Therefore, the increase in particle size (from 200 nm to 500–1000 nm) and in membrane fluorescence (10 a.u.–150 a.u.; Fig.6B,D) and the shift between the two populations suggests clustering of 15 or more vesicles to form larger particles. The shift

from single vesicles (orange) to larger particles (dark cyan) occurred in a hGBP1fn concentration-dependent manner. Analysis of the number of detected events in the two populations showed that a steady increase of the population of tethered vesicles takes place in the range of 5–10 µ^M hGBP1_fn resulting in a constant number of tethered vesicles for 10–15 µ^M hGBP1_fn (Fig. 6G).

Since GDP*AlF_X mimics the transition state of the first cleavage step in GTP hydrolysis, hGBP1_fn is kept irreversibly in a stable conformation having this nucleotide bound in its binding pocket, making also the effect of tethering static and irreversible. To con- sider a more dynamic system for investigating the tethering effect of hGBP1_fn, the natural nucleotide GTP was offered as substrate to induce membrane binding and liposome tethering. In the presence of GTP, hGBP1fn establishes an equilibrium of membrane bound and soluble protein, which is dependent on GTP hydrolysis. The hGBP1fn-induced tethering effect was observed for two different protein concentrations (10 µ^Mand 15µ^M) over time (Fig. 6H,I). Protein concentrations were chosen, as tethering was only observed for hGBP1_fn concentration≥7.5µ^M. The number of events in the two different populations was evaluated and demonstrated the reversibility of the tethering effect in the presence of GTP. For both tested concentrations, the amount of tethered vesicles in the second population increased over time reaching a maximum after 25 min (Fig. 6H,I: dark cyan). After that, the vesicle population shifted back, which increased again the number of single vesicles in the first population (Fig.6H,I: orange). Here, it is important to note that the decrease in the second population

Fig. 6.hGBP1fn binding to liposomes induces vesicle tethering. Large unilamellar vesicles (LUVs) were incubated with the indicated concentrations of Alexa488-labeled hGBP1_fnin the presence of GDP*AlF_Xor GTP and analyzed by flow cytometry. (A, C) Representative dot plots (sideward scatter vs. hGBP1_fn-derived fluorescence) of three replicates. The respective regions applied for data analysis are depicted in orange (population 1) and dark cyan (population 2). (B, D) Sideward scatter (SSC) and membrane derived fluorescence (vesicles composed of BPL and 0.5 % ATTO633-labeled DOPE) in the presence of 0.5µ^M (B) or 12.5µ^M (D) hGBP1fn and 250µ^M GDP*AlFX. The two populations used for further analysis are depicted in orange (population 1, single vesicles) and dark cyan (population 2, tethered vesicles).

The 0.2µm (red), 0.5µm (green), 1µm (blue), and 2µm (cyan) boxes correspond to the size gates defined by the measurement of beads with known size (F). E: Forward scatter (FSC) and sideward scatter (SSC) of the LUVs (black) in the presence of 12.5µ^M hGBP1fnand 250µ^M GDP*AlF_X. The two populations used for further analysis are depicted in orange (population 1, single vesicles) and dark cyan (population 2, tethered vesicles) and were chosen to represent two distinct populations. F: Fluorescent beads of defined sizes (diameter:

0.2µm (red), 0.5µm (green), 1µm (blue), and 2µm (cyan); impurities in gray) in the sideward scatter and the bead derived fluorescence.

The differently sized beads define distinct regions in the sideward scatter that can be applied on the size analysis of the LUVs. G: Evolution of the vesicle number in population 1 (orange) and population 2 (dark cyan) in the presence of the indicated hGBP1_fnconcentrations. The mean value (SD) of three replicates is shown. H, I: Time-dependent evolution of the vesicle number in population 1 (orange) and population 2 (dark cyan) in the presence of 10µ^M(H) or 15µ^M(I) hGBP1_fnbound to the LUVs in a GTP-dependent manner. A representative measurement out of three replicates is shown. J: Titration curve of GDP*AlF_X-induced binding of hGBP1_fnto LUVs. The geometrical mean fluorescence of population 1 is plotted, and a linear fit is applied to the increase in fluorescence intensity (red line) and the saturation of bound protein (gray line). The mean value (SD) of three replicates is shown.

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occurred faster for 15µ^M than for 10µ^M hGBP1_fn, which is explained by the faster depletion of GTP by hydrolysis at higher hGBP1 concentration and therefore earlier detachment from the membranes [25,54,55].

Next, we asked in what orientation the hGBP1 molecules are arranged on the membrane and we used flow cytometry analysis to address the density of hGBP1_fn molecules on the membrane surface. There- fore, the protein-derived fluorescence intensity (geometrical mean fluorescence) in the first population (orange) was further evaluated (Fig.6J). For low concentrations of hGBP1_fn (0–5 µ^M), the protein fluorescence intensity on the vesicles increased linearly with increasing protein concentration (red line, slope: 3.8 a.u./µ^M). At a concentration of 8 µ^M hGBP1_fn, the protein fluorescence on the vesicles reached saturation (gray line), suggesting complete coverage or saturation of the vesicle surface. Interestingly, the onset of the saturation in protein fluorescence coincides with the shift of the vesicle population from single to clustered vesicles (Fig.6G,J), suggesting that only completely protein covered vesicles are tethered to larger aggre- gates. Furthermore, the knowledge of the concentration at which hGBP1 saturates the vesicle surface allowed us to predict what orientation the hGBP1 molecules adapt on the membranes. We considered two possible scenarios: The hGBP1 molecule with the farnesyl tail integrated into the membrane can either lay alongside (‘flat’) onto the membrane surface adapting a closed conformation which allows the formation of head-to-head dimers or it adapts an open conformation with the LG domain pointing outwards alongside with other open hGBP1 molecules forming additional MD:MD contacts as suggested above. In the first case, given the dimensions of the protein molecule, hGBP1 will have a footprint on the membrane surface of 60 nm². In the second case, the footprint is estimated to be 20 nm² based on the dimensions of the LG domain (PDB: 2D4H, PyMOL). The head group of a single lipid molecule is estimated to be 0.7 nm². A comparison of the foot- prints reveals lipid/hGBP1 ratios of 85.7 for hGBP1 laying in a closed conformation on the membrane and 28.6 for hGBP1 adapting the outstretched conformation. The molar ratio of hGBP1 saturating the vesicle surface (8µ^M hGBP1) and the lipid molecules in the outer lipid layer (250µ^M lipid) equals 31.3, which approximately matches the footprint ratio of 28.6, revealing that hGBP1 molecules are arranged in a parallel fashion with their farnesyl tails anchored in the membrane and their LG domains pointing outwards.

Discussion

In our earlier study, we showed that the farnesyl tail attached to the C-terminal cysteine of hGBP1 is crucial for nucleotide-dependent membrane binding of the protein as well as for the formation of structured hGBP1 polymers [38]. We demonstrated in the same study that this hydrophobic moiety contributes to the stabilization of the closed structure of hGBP1 [38].

This finding was supported recently by Ji et al. [43]

who solved the X-ray structure of farnesylated hGBP1 and revealed that the farnesyl tail resides in a hydrophobic groove established by the middle and GTPase effector domain of the protein. The release of the farnesyl tail from this hydrophobic pocket is nucleotide controlled and although GTP binding can trigger membrane binding GTP hydrolysis is mandatory for polymerization[38].

Results presented in this study revealed that GTP hydrolysis is also relevant for membrane binding (Fig. 2). Addition of GTP results in more and GDP*AlF_X even much more hGBP1 binding to the GUV as compared to the nonhydrolysable GTPcS.

These results demonstrate that GTP hydrolysis is not only mandatory for polymerization but also induces or stabilizes a state that increases hGBP1’s ability to attach to membranes. This implies that similar structural changes in the protein might be responsible for both processes. Possibly, a shift of the structural equilibrium toward the open state is stabilized by MD:MD contacts between two hGBP1 molecules as we could rationalize for polymerization [44]. This scenario would explain a persistent exposure of the farnesyl tail.

The more or less strong attachment of hGBP1 to the GUV as we observe it for different nucleotides and nucleotide analogs can be explained by the nucleotide- dependent farnesyl switch. The understanding of the farnesyl switch in the sense of opening a structure more or less widely does not make sense as such structural changes will occur cooperatively. Rather the equilibrium between an off and on state of a switch is shifted to a different extent by the various nucleotides.

Only a small fraction of hGBP1_fn is opened in the presence of GTPcS for instance and a large fraction or almost all is opened when hGBP1_fn is bound to GDP*AlFX. Our previous investigations on the mechanism of hGBP1fn polymerization [44] led us to the conclusion that the shift of the equilibrium between the closed and open structure is not only a matter of nucleotide binding and intramolecular changes of the structure. Rather the open structure is stabilized by dimer formation of the protein and mutual stabilization of the open state.

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To test whether this scenario is also important for membrane binding, we challenged here the nucleotide- dependent behavior of the RK mutant, which is more prone to opening due to the weakened contacts between the GED and LG/MD. For this mutant, the equilibrium is still on the closed conformation side when no nucleotide is present or when GMP is present as no membrane attachment was observed in the GUV binding assay (Fig.3). This demonstrates that in the monomeric state (apo, GMP bound) the hydrophobic interaction of the farnesyl moiety with the residing pocket is intact keeping the structure closed, independent from the electrostatic attraction between the LG domain and GED. The addition of GDP or the transition state analog GMP*AlF_X enables membrane attachment of the RK mutant to a higher extent as for the wild-type, since the RK mutant reveals a signifi- cant higher activity on the hydrolysis of GDP compared to wild-type hGBP1 [27,33]. The higher extent of binding was defined by the relative number of protein-bound vesicles (Fig.3), whereby the observation of either protein-bound or empty vesicles already pointed toward a stabilization of membrane-bound protein by network formation.

As we know that GTPase activity of hGBP1 as well as GDPase activity of a mutant like RK is intimately coupled to dimerization through the LG domains we addressed the issue of LG:LG contact formation.

Firstly, the comparison between 2µ^M and 10µ^M of hGBP1-RK shows an increase in membrane binding, which is much more than an expected linear increase.

This is explained by the shift of the equilibrium from monomers to dimers at higher concentrations. Sec- ondly, and more directly, we challenged the importance of dimerization by utilizing the point mutant R244A which lacks one of the arginines stabilizing the interface of the LG:LG dimer. In fact, this mutant failed to attach to the GUV in the presence of GTP and only a 10-fold increased protein concentration shifting the equilibrium toward dimers showed weak membrane attachment (Fig.3). Thus, dimerization is absolutely crucial to establish a protein conformation in which a proper position of the farnesyl moiety is stabilized in order to bind to membranes.

In fact, we were able to demonstrate that for membrane attachment it is a matter of farnesyl positioning by dimerization rather than increasing the number of farnesyl moieties. As shown by mixed dimer formation, only one farnesyl moiety is required to reach membrane attachment (Fig.5). More intriguingly, using the deletion mutants hGBP1-DGED and hGBP1-DMD/GED, which corresponds to hGBP1- LG, in the mixed dimer set-up, we demonstrated that

the MD domains are necessary for stabilization of the open state. Combining the results from the GUV binding assay and intermolecular FRET measurements, we suggest that dimerization through the LG domains is not sufficient to expose the farnesyl tail persistently but that at least the middle domains have to be present in order to enable hGBP1_fn for membrane attachment.

Despite the different dependencies of polymerization and membrane attachment on nucleotide binding and hydrolysis, our conclusions regarding the structural requirements for the two processes approach each other. In the case of polymerization, we suggested a model in which the GTPase activity induces dimerization and opening. Then, mutual stabilization of the open form through MD:MD contacts leads to the establishment of a structured polymer where the open hGBP1 molecules are aligned alongside to each other.

Here, we come to a similar scenario as pointed out above. In addition, the stable protein coat in the presence of GDP*AlF_X around the vesicle (Fig. 4), which even persists after removal of the lipid bilayer reminds strongly of the polymers stabilized by GDP*AlF_X in our previous work [44]. The difference in behavior of the protein coats induced by GTP and by GDP*AlF_X, respectively, might be based on the dynamics of the systems. GTP is hydrolyzed by hGBP1_fn, and even if the specific turnover number is reduced in the presence of membranes, the proteins are in a dynamic exchange between its soluble and membrane-bound form by hydrolyzing GTP [38]. In contrast, GDP*AlFX is known to induce strong and irreversible interactions [44], producing a stable protein coat, suggesting that the hGBP1fn molecules are able to form a tight polymer network on the membrane.

Utilizing flow cytometry, we made two interesting observations (Fig.6): Firstly, the concentration of hGBP1 on the surface of lipid vesicles reached a saturation value when the molar ratio of outer surface lipid:hGBP1 reached a value of roughly 30:1. Taking into account the ratio of footprint areas of a lipid head group and the LG domain of hGBP1, which equals almost 1:29, we can conclude that the membrane outer surface area can accommodate a monolayer of hGBP1 molecules supposed the open structure of hGBP1 stands upright on the membrane surface like pins in a pincushion (in contrast to the scenario when the hGBP1 molecules are lying ‘flat’ on the surface the footprint ratio equals 1:86). Secondly, the hGBP1-induced tethering led to an agglomeration of roughly 15 vesicles as judged from the increase in particle size and membrane fluorescence. Intriguingly, tethering was only observed after a hGBP1

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concentration of 5–8µ^M was reached which we identi- fied above as the concentration required to completely cover the membrane surface.

Taken together, our observations on hGBP1´s nucleotide-dependent membrane binding, tethering and surface saturation culminate in a model which—not surprisingly—shows similar features as the one for polymerization (Fig.7). GTP binding shifts the equilibrium from the closed to the open structure, which implies the abolishment of the farnesyl tail:MD/GED contact. The dimerization of the protein concomitant with the opening allows MD:MD contact formation which we suggest to further stabilize the open structure and thereby a persistent exposure of the farnesyl groups available now for interaction with each other in case of polymerization or for membrane insertion.

In case of the latter, an upright orientation of the open hGBP1 molecules on the membrane surface becomes more, and more likely, the more hGBP1 cover the surface as the orientation of hGBP1 molecules alongside to each other is favored by MD:MD as well as LG:

LG contacts. This pins in a pincushion scenario reached at saturation enables the hGBP1 covered vesicles to tether as the LG domains are located in the outer sphere ready to interact with LG domains from a neighboring vesicle (Fig.7).

Our conclusions as to the nucleotide triggered structural changes of hGBP1 and its assembly on the artificial membrane surface will help future studies to understand the mechanism of hGBP1 interactions with host and pathogen membranes.

Materials and methods

Protein expression and purification

Protein expression, farnesylation, and purification were performed as described earlier[30,44,56]. In short, hGBP1 and the different mutants were cloned into pQE80L and expressed in BL21-CodonPlus (DE3)-RIL cells (Stratagene, Heidelberg, Germany). Protein was purified via affinity chromatography (HisPur Cobalt Resin, Thermo Fisher Sci- entific, Waltham, MA, USA) and size-exclusion chromatography (SEC) (Superdex 200 26/60, 320 mL, GE Healthcare, Munich, Germany). hGBP1 and hGBP1-Q577C were farnesylated by enzymatic modification, and the mutants hGBP1-RK and hGBP1-R244A were co-expressed with recombinant FTase as described in [39,56]. Farnesylated proteins were additionally purified via hydrophobic interaction chromatography (Butyl Sepharose High Performance Column, 20 mL, GE Healthcare) before SEC[56].

Labeling of protein with fluorescent dye

For labeling of the different hGBP variants, AlexaFluo- r488-C5- and AlexaFuor647-C2-maleimide dyes (Thermo Fisher Scientific) were used. In general, coupling of fluorescent dye to accessible cysteines of hGBP1 were performed on ice under exclusion of light in buffer L (50 mM Tris/

HCl pH 7.4, 5 mM MgCl₂, 150 mM NaCl). For labeling with Alexa Fluor488-C5- maleimide, hGBP1fn, hGBP1, hGBP1-ΔGED, hGBP1-R244Afn, and hGBP1-RKfn were incubated in an equimolar ratio with the fluorescent dye for 15 min (farnesylated proteins), 10 min (hGBP1) or 30 min (hGBP1-ΔGED). hGBP1-LG was labeled for 60 min with AlexaFluor488-C5-maleimide using a molar ratio of 2.0 dye/protein. Finally, hGBP1fnwas labeled with AlexaFluor647-C2-maleimide for 20 min using a molar ration of 1.0 dye/protein. All labeling reactions were stopped by the addition of 2 mMdithiothreitol (DTT) and a)

b) c)

d) e) f)

RK mutant GTPase

GMP

tethering

hGBP:lipids

1:30 15 LUVs

hGBP1-loaded LUV

Fig. 7.hGBP1’s requirements for membrane binding and tethering effect of hGBP1 bound to membranes. The upper panel of the scheme illustrates the equilibrium between the closed and the open state. A shift of the equilibrium to the open state is favored a) in the presence of GTP, b) by weakening the contact between GTPase domain and C-terminal domain, and c) by the support of a hGBP1 dimer partner which harbors at least the GTPase domain and the middle domain. The closed state of hGBP1 is stabilized d) in the absence of nucleotide or in the presence of GMP, e) by the farnesyl tail residing in its hydrophobic pocket, and f) by the formation of a dimer with a hGBP1 partner which lacks the middle domain. The lower panel shows a vesicle completely covered by hGBP1 in the open state (not to scale, hGBP1 has a length of 30 nm and a vesicle has a diameter of 200 nm). These hGBP1 loaded vesicles are tethered through contacts between the surface exposed GTPase domains giving rise to an aggregate of 15 or more vesicles in size. Color code as in Fig.1.

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unbound dye was removed by buffer exchange via ultrafiltration (Vivaspin Turbo 4; 10 kDa cutoff, Sartorius, G€ottingen, Germany) to buffer C (50 mMTris/HCl pH 7.9, 5 mM MgCl₂, 150 mM NaCl). Afterward, the protein was concentrated by ultrafiltration. Protein concentrations and labeling efficiencies were calculated according to Lambert- Beer law from absorption intensities at wavelengths 280 nm (hGBP1), 491 nm (Alexa488, donor) and 651 nm (Alex- a647, acceptor) in buffer C supplemented with 2 mMDTT, using molar absorption coefficients ofehGBP1-FL: 45440 M^-

1cm^-1, ehGBP1-ΔGED: 44,350 M^-1cm^-1, ehGBP1-LG: 35,410 M^-

1cm^-1,e491: 71,000 M^-1cm^-1,e647: 268,000 M^-1cm^-1, and the correction factors provided by the company. Labeling efficiencies were as follows: hGBP1_fn: LE₄₈₈: 30 % and 85%

(used for flow cytometry), LE₆₄₇: 20 %; hGBP1: LE₄₈₈: 90 %; hGBP1-ΔGED: LE488: 46 %; hGBP1-R244Afn: LE488: 48 %; hGBP1-RKfn: LE488: 25 %; hGBP1-LG:

LE488: 111 %.

Liposome preparation

All lipid stocks were dissolved in chloroform and stored at 20°C. Giant unilamellar vesicles (GUVs) were prepared by electroformation using Pt wires as electrodes as described previously [42]. Porcine brain polar lipids (BPL, Avanti Polar Lipids, Alabama, USA) and 1,2-dipalmitoyl- sn-glycero-3-phosphoethanolamine (Avanti Polar Lipids) were mixed in chloroform (to final concentrations of 2 mg•ml^-1 and 40µg•ml^-1, respectively) and applied to Pt wires. Chloroform was removed under vacuum, and the wires were sealed with a Teflon cap filled with sucrose solution. The sucrose solution was adjusted to the osmolarity of buffer C. GUVs were electroformed by applying a sinu- soidal voltage for 4 h (1.3 V, 12 Hz). Detachment of the GUVs from the wires occurred by applying a second sinu- soidal voltage for 30 min (2.0 V, 4 Hz). GUVs were stored on ice after electroformation and always used within 1-5 h after electroformation.

Large unilamellar vesicles (LUVs) were prepared from BPLs supplemented with 0.5 mol% ATTO633-labeled 1,2- dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE) (ATTOTEC, Siegen, Germany) to allow detection of liposomes by flow cytometry. A homogenous lipid film was generated in a glass tube under nitrogen flow, and residual chloroform was removed under vacuum for 30 min. Rehy- dration buffer (bufferR: 50 mM Tris/HCl, 5 mM MgCl2, 150 mM NaCl, 10 % w/v sucrose, pH 7.9) was added to reach a final lipid concentration of 3 mg•ml^-1. The lipid film was rehydrated in a water bath at 55°C for 30 min.

After four freeze (liquid nitrogen) and thaw (55°C, water bath) cycles, size extrusion using a pore size of 800 nm (polycarbonate membrane, Whatman, Maidstone, UK) and a mini extruder (Avanti Polar Lipids) was performed at 55°C. Vesicles were stored at 4°C and used the same day.

GUV binding assay and confocal microscopy

Binding of hGBP1 to GUVs was examined using a Leica Microsystems TCS SP8 confocal laser scanning microscope (Wetzlar, Germany) equipped with a 63x/1.2 NA water immersion objective under temperature control (25°C).

Self-assembled chambers ([57], placed on cover slips #1.5, Thermo Scientific) were coated with 5 mg•ml^-1 BSA for 10 min and washed twice with buffer C. The chamber was filled with 100µL buffer C (or buffer C+AlF_x: 300µ^M AlCl3, 10 m^M NaF) supplemented with 50µ^M BSA and 2µL GUVs were added. If not indicated otherwise, 2µ^M Alexa488-labeled hGBP1 was mixed with nucleotide (250µ^M GMP, GDP*AlFX, GMP*AlFX, 500µ^M GTPcS, or 2 mM GTP, GDP) in 50µL buffer C (or buffer C+AlFx) and added to the GUVs immediately (final concentrations given here for protein and nucleotide for the final volume of 150µL). For the delivery of truncated hGBP1fn, 2 µ^M hGBP1fnlabeled with Alexa647 was mixed with either 2µ^M hGBP1 or 5µ^M hGBP1-DGED or hGBP1-LG (hGBP1, hGBP1-DGED, and hGBP1-LG, all labeled with Alexa488).

To solubilize the vesicles after binding of hGBP1_fn, 0.1%

(v/v) Triton X-100 was added to the sample chamber and time-lapse imaging was started. Images were collected every 2–7 s using Leica Imaging Software (LasX). The following excitation wavelengths, laser powers, and spectral detection windows on the photomultiplier (PM; gain 700 mV) and hybrid detectors (HyD; gain 20–40%) were used: 488 nm, 10–30%, HyD, 500–550 nm (Alexa488); 651 nm, 10%, HyD, 660–730 nm (Alexa647); 560 nm, 1–3%, PM, 590– 640 nm (Rhodamine). Measurements in which fluorescence intensities were compared quantitatively (Fig.2A,B and Fig.3D,E GTP) were performed under constant microscope settings, respectively. For the other measurements, laser power and detector gain were chosen and changed to obtain a reasonable fluorescence signal.

The measurements displayed in Fig.2B were performed on a confocal fluorescence microscope (Nikon) using two laser lines with excitation wavelength of 488 nm (Alexa488) and 561 nm (Rhodamine). Emission was detected after passing respective band pass filters (525/50 and 595/50). Z- stacks of vesicles were taken with identical laser power and gain settings.

Intermolecular FRET measurements

Intermolecular FRET measurements were performed on an LS55 fluorescence spectrometer (Perkin Elmer, Waltham, MA, USA). In general, the donor fluorophore (Alexa488) was excited at 498 nm and the acceptor fluorescence (Alex- a647) was detected at 664 nm. Excitation and emission slits were set to 10 nm and 15 nm, respectively, and detector voltage was set to 775 mV. 2µ^MAlexa647-labeled hGBP1_fn was mixed with either 2µ^M hGBP1, 5µ^M hGBP1-ΔGED,

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or 5µ^M hGBP1-LG (all labeled with Alexa488) in buffer C+ AlFx (supplemented with 50µ^M BSA). Interac- tion was induced by addition of 250µ^M GDP at t=0 s.

Measurements were carried out at 25°C. Changes in fluorescence at 664 nm (F) were normalized by initial fluorescence (F0) prior to addition of nucleotide (plotted as F/F0).

LUV binding assay

To determine the nucleotide induced binding of hGBP1fnto LUVs, the protein was labeled with Alexa488. Increasing concentrations of Alexa488 labeled hGBP1fn (0.5–15µ^M) were incubated with 0.5 mg•ml^-1vesicles in bufferRsupplemented with 10 mM NaF, 300µ^M AlCl3 (for GDP*AlFX-induced binding), and 0.05 % (w/v) BSA. Binding was induced by addition of either 3 mMGTP or 250µ^MGDP*AlF_X, respectively. Samples with GDP*AlF_Xwere analyzed 1 min after nucleotide addition by flow cytometry. For the time-lapse measurements of GTP-induced binding, samples with either 10µ^M or 15µ^M hGBP1fn were prepared with 0.5 mg•ml^-1 vesicles in buffer_Rcontaining 0.05 % (w/v) BSA. The first measurement was performed after 1 min. After that, the same sample was measured every 5 min for 60 min.

Flow cytometry

Flow cytometry was performed on a Becton Dickinson FACS Calibur (San Jose, CA) equipped with an 488-nm argon laser and 635-nm red-diode laser using Cell Quest software. Size calibration on the forward and sideward scatter was com- pleted with fluorescent beads of defined sizes (0.2–2µm, Thermo Scientific). To track the hGBP1fn derived fluorescence, Alexa488 was excited by the argon-ion laser (15 mW, kexc: 488 nm) and emission acquired in FL1-H (PMT, 530/

30 nm). ATTO633-labeled LUVs (0.5 mgmL^-1) were excited by the red-diode laser (10 mW,kexc: 635 nm) and emission acquired in FL4-H (PMT, 661/16 nm). Thirty thousand events were analyzed without gating during acquisition.

Data analysis

For the GUV binding assay, confocal images were processed with Fiji ([58], https://imagej.net/Fiji) as described previously [38]. The plotted fluorescence values represent the protein fluorescence on the membrane corrected for the fluorescence derived from unbound protein in the background (F=FmembraneFbackground). Membrane fluorescence was also considered with respect to the background in the rhodamine channel (dissolution of the membrane, Fig.4).

For the LUV binding assay, data were processed with Flowing Software 2.5.1 (Cell Imaging Core, Turku Centre Biotechnology) based on Temmerman et al.[59]. Vesicles were selected based on forward and side-scatter gating and two different regions were defined, representing single and aggregated

vesicles, respectively. The number of events and the fluorescence intensity in the green and red channel were acquired in the determined regions, respectively. Subsequently, histograms of the green fluorescence (Alexa488) and the red fluorescence (ATTO633) were generated to calculate the geometrical mean fluorescence intensity of the total liposome population. Analy- sis was performed for>5000 events per population. Maxi- mum binding of hGBP1fn to the vesicles was obtained by plotting the geometrical mean fluorescence of the first region against the respective protein concentration. The comparison of the number of events in each region for each protein concentration was used to demonstrate the aggregation of the vesicle population.

To observe GTP-dependent vesicle tethering over time, the same gate as used for GDP*AlFX-induced binding was applied to the hGBP1fn-derived fluorescence of each measured time point, to obtain the number of events in the different regions.

Acknowledgement

Supporting grants were from the German Research Foundation INST 213/886-1 FUGG to T.G.P and HE2679/6-1 to CH, and Project International (PR.INT) Fellowship via RUB Research School PLUS [DFG GSC 98/3] and DFG Research Fellowship 427472513 to MK.

Author contributions

LS planned and performed experiments, analyzed data, discussed results, and wrote the paper. LDS performed experiments, analyzed data, and discussed results. MK planned and performed experiments, analyzed data, discussed results, and reviewed and edited manuscript.

AR planned experiments. TGP planned experiments, analyzed data, discussed results, and reviewed and edited manuscript. CH planned experiments, analyzed data, discussed results, and wrote paper.

Conflict of Interest

The authors declare no conflict of interest.

Peer Review

The peer review history for this article is available at https://publons.com/publon/10.1111/febs.15703.

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