Prolonged display or rapid internalization of the IgG-binding protein ZZ anchored to the surface of cells using the diphtheria toxin T domain

Philippe Nizard1, Alexandre Chenal1, Bruno Beaumelle2, Alain Fourcade3 and Daniel Gillet1,4

1 Département d'Ingénierie et d'Etudes des Protéines and 3 Département de Biologie Cellulaire et Moleculaire, CEA–Saclay, 91191 Gif sur Yvette cedex and 2 UMR 5539 CNRS, Université Montpellier II, 34095 Montpellier cedex 05, France


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We have shown previously that the diphtheria toxin transmembrane domain (T) may function as a membrane anchor for soluble proteins fused at its C-terminus. Binding to membranes is triggered by acidic pH. Here, we further characterized this anchoring device. Soluble proteins may be fused at the N-terminus of the T domain or at both extremities, without modifying its membrane binding properties. This allows one to choose the orientation of the protein to be attached to the membrane. Maximum binding to the cell surface is reached within 1 h. Anchoring occurs on cells previously treated with proteinase K, suggesting that T interacts with the lipid phase of the membrane without the help of cell surface proteins. Binding does not permeabilize cells or affect cell viability, despite the fact that it permeabilizes liposomes and alters their structure. When attached to L929 fibroblasts, the proteins are not internalized and remain displayed at their surface for more than 24 h. When bound to K562 myeloid cells, the molecules are internalized and degraded. Thus, depending on the cell type, soluble proteins may be anchored to the surface of cells by the T domain for an extended time or directed towards an internalization pathway.

Keywords: diphtheria toxin/immunoglobulin/membrane anchor/protein A/transmembrane domain


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cell engineering technologies may offer new approaches for the treatment of various diseases, including cancer and degenerative or genetic diseases. The aim is to constrain the cell to secrete or express at its surface a protein, which will modify its physiology or its relations with neighboring cells. Most of the time, the manipulation of the cell is based on the transfer of genetic material. However, the establishment of genetically modified cells is cumbersome in a clinical context and suffers from several drawbacks. Some cells are difficult to transfect. The selection of stable transfectants may take several days. The injection of cells genetically modified with the help of viruses may be hazardous (Marshall, 1999Go). The development of totally safe viruses has not yet been accomplished (Alemany et al.2000Go), as viral recombination or adverse immune reaction may occur (Simon et al.1993Go). Finally, the amount of recombinant proteins produced by transfected cells is difficult to control (Schmidt et al.1995Go), requiring the selection of clones or the use of tunable promoters and the measure of recombinant gene expression.

Our goal is to develop an alternative strategy to gene transfer, to anchor a given protein at the surface of cells. This strategy is based on the development of a protein membrane anchor derived from the diphtheria toxin transmembrane (T) domain (Liger et al.1998Go; Nizard et al.1998Go).

Diphtheria toxin is a 58 kDa protein organized in three domains: receptor binding (R), transmembrane (T) and catalytic (C) (Bennett and Eisenberg, 1994Go). After binding to its cell surface receptor, the toxin is internalized. The acidic pH in the endosome triggers the insertion of T in the membrane. This assists the translocation of C to the cytoplasm where it blocks protein translation by ADP-ribosylation of elongation factor 2 (Lemichez et al.1997Go). The structure of the T domain (about 22 kDa) is organized in three layers of {alpha}-helices (Bennett and Eisenberg, 1994Go): a central hydrophobic helical hairpin sandwiched and hidden from the solvent by two amphiphilic layers. Acidic pH induces a conformational change leading to exposure of the hydrophobic parts of the central layer, promoting interaction with the membrane (Zhan et al.1995Go).

We have shown previously that the T domain may function as a membrane anchor to attach soluble proteins fused to its C-terminus on the surface of cells. This was done for cytokines (Liger et al.1998Go) and the IgG binding protein ZZ (Nizard et al.1998Go). ZZ was generated by duplication of a mutated B domain from the staphylococcal protein A (Nilsson et al.1987Go; Ljungberg et al.1993Go; Jansson et al.1998Go). The fusion protein T-ZZ, once bound to the surface of lipid vesicles or cells, was able to bind IgG, thus functioning as a membrane anchor for antibodies. Anchoring to membranes is fast and easily triggered by incubation at a mildly acidic pH (pH 5).

In the present work, we further characterized the properties of the T domain as a membrane anchor. Using the ZZ protein as a model, we studied the possibility of fusing proteins to its N-terminus or to both extremities. This was intended to establish whether it is possible to choose the orientation of the protein to be anchored, its N- or C-terminus towards the membrane. We compared the membrane binding properties of the different constructs. Finally, we investigated the fate of the molecules once attached to the surface of cells.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Expression vectors

Vector pCD2-ZZ for the expression of the fusion protein T-ZZ (Nizard et al.1998Go) and vector pCP for the expression of the protein ZZ (Drevet et al.1997Go) have been described previously. All other plasmids were derived from those. pA/T encoding the isolated T domain was obtained, first by the digestion of pCD2-ZZ with SphI and HindIII to remove the sequence encoding ZZ and then by cloning in place a DNA fragment made by hybridization of the oligonucleotides 5'-CTT AGT AAA-3' and 5'-AGC TTT TAC TAA GCA TG-3' in order to introduce two stop codons at the end of the T domain coding sequence. pA/ZZ-T encoding the fusion protein ZZ-T was constructed, first by the digestion of pCP with XmaI and BamHI to remove the 3' multiple cloning site and then by cloning in place the T coding fragment synthesized by PCR amplification from pCD2-ZZ using the primers 5'-CGG GGG TTC TGG TGG TTC TGG AGG TTC TGG TGG TTC TGG GTC TGG TTG CAT CAA CCT GGA TTG GGA-3' and 5'-GCA GCT GGA TCC TTA TCA AGC ATG CGT CTT GTG ACC C-3'. These primers were designed to introduce the restriction sites XmaI and BamHI for cloning and a sequence encoding a hydrophilic peptide spacer between ZZ and T (Table IGo). Finally, pZZ-T-ZZ encoding the triple fusion protein ZZ-T-ZZ was constructed by digestion of both plasmids pA/ZZ-T and pCD2-ZZ with XbaI and ClaI located within the promoter region of the plasmids and in the T domain coding region, respectively. The fragment encoding ZZ and half of T from pA/ZZ-T was then cloned in the digested pCD2-ZZ vector carrying the remaining T coding region followed by a ZZ coding sequence.


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Table I. Primary structure of the recombinant proteins
 
Protein expression and extraction

Expression was done in BL21(DE3) cells at 37°C. Bacteria were grown in Terrific Broth (12 g/l tryptone, 24 g/l yeast extract (Difco, Detroit, MI), 0.4% (v/v) glycerol, 17 mM KH2PO4, 72 mM K2HPO4) for proteins ZZ, ZZ-T and ZZ-T-ZZ and in M9 [5 mM (NH4)2SO4, 22 mM KH2PO4, 39 mM Na2HPO4, 39 mM NaH2PO4, 8.5 mM NaCl, 1mM MgSO4, 0.5 µM FeCl2, 0.1 mM CaCl2, 5 g/l glucose] for proteins T and T-ZZ. All media were supplemented with 0.2 mg/l ampicillin and pH was kept around 7.2 during culture. When cultures reached mid-logarithmic phase (OD600 = 0.6–0.7 ), protein expression was induced with 0.1 mM IPTG (Fluka, Buchs, Switzerland) carried out for 4 h. Cells were harvested by centrifugation at 4500 g for 30 min at 4°C and resuspended in 15 ml of TN buffer (50 mM Tris–HCl, pH 7.8, 150 mM NaCl), excepted cells transfected with plasmid pA/T that were resuspended in 15 ml of PNI-10 buffer (20 mM sodium phosphate, pH 7.8, 0.5 M NaCl, 10 mM imidazole). After re-suspension, cells were supplemented with 0.1 ml of 0.1 M PMSF (Sigma) in ethanol and 1 ml of a 10 mg/ml solution of lysozyme (Sigma). After incubation for 1 h on ice, extracts were homogenized by repeated passage through a needle with a syringe followed by sonication for 10 min (power setting 60%, pulse 1 s, rest 1 s). The soluble cytoplasmic fraction was recovered by centrifugation at 18 000 g for 30 min at 4°C. The pellet was re-suspended in the same buffer, sonicated again and the second soluble fraction was recovered by centrifugation. Both soluble fractions were pooled and treated with 1 µg/ml of DNase I and RNase A for 1 h at 24°C and filtered through 0.45 and 0.22 µm filters. Before purification, soluble fractions were kept on ice to avoid protein degradation and aggregation.

Purification of the recombinant T domain

The T domain was purified by immobilized metal ion (Ni+) affinity chromatography (IMAC) using a HiTrap Chelating Sepharose Fast Flow gel (Pharmacia, Uppsala, Sweden). A 5 ml volume of gel was prepared according to the manufacturer's instructions. The protein sample was loaded and the column was successively washed with PNI-50 and PNI-70 buffers (20 mM sodium phosphate, pH 7.8, 0.5 M NaCl, 50 and 70 mM imidazole). Elution was done with PNI-500 buffer (20 mM sodium phosphate, pH 7.8, 0.5 M NaCl, 0.5 M imidazole). The eluted proteins were treated with 14.3 mM 2-mercaptoethanol for 1 h at 24°C to impair any intermolecular cystine formation. Then, imidazole, NaCl and 2-mercaptoethanol were removed by buffer exchange with 20 mM sodium phosphate, pH 7.8 on a Sephadex G-25 Superfine Hitrap desalting column (Pharmacia Biotech, Uppsala, Sweden). All proteins were stored in 20 mM sodium phosphate, pH 7.8 at –20°C.

Purification of the recombinant proteins ZZ, T-ZZ, ZZ-T and ZZ-T-ZZ

Purifications were conducted by affinity chromatography on an IgG Sepharose 6 Fast Flow column according to the manufacturer's protocol (Pharmacia Biotech), except for the washing buffer, the pH of which was adjusted to 4.5 instead of 5. The eluted proteins were lyophilized and stored at –20°C as such or after re-suspension in 20 mM sodium phosphate, pH 7.8.

Lipid vesicles permeation study

Large unilamellar vesicles (LUV) were prepared as described for the atomic force microscopy study below and the permeabilization study was carried out as described previously (Nizard et al.1998Go).

Atomic force microscopy study

LUV in PBS were prepared by reversed-phase evaporation as described (Rigaud et al.1983Go) using a mixture of egg phosphatidylcholine and egg phosphatidic acid (Avanti Polar Lipids, Alabaster, AL) at a molar ratio of 9:1. LUV at concentration of 2 mg/ml (2.6 mM lipids) were incubated with the fusion protein T-ZZ at a concentration of 50 µg/ml (1.3 µM) in PBS supplemented with 10 mM acetate, at a final pH of 4.8 or 7.4, for 1 h at 20°C. In controls, LUV were incubated at both pH in the absence of protein. Preparations were then deposited on mica slabs to allow the LUV to bind to the mica surface at room temperature. The slabs were then gently rinsed with water to remove unbound material and salts, which interfere with atomic force microscopy scanning, and allowed to dry. Samples were observed by scanning of 4x4 or 7.5x7.5 µm areas in non-contact mode with the tip of a Topometrix microscope.

Cell surface binding immunodetection assay

Assays on adherent L929 cells were performed in 96-well plates as described previously (Liger et al.1998Go; Nizard et al.1998Go), except that vizualization was done as described below for the assay on A20 cells. Assays on A20 cells were done on suspensions of 6x106 cells in 15 ml Falcon tubes and all washes were done by centrifugation at 1200 r.p.m. in PBS pH 7.4. After three washes, the recombinant proteins were added to the cells at the indicated concentration in PBS supplemented with 10 mM acetate at pH 7.4 or 4.8. After incubation for 1 h at 20°C, cells were washed once and transferred to a new tube. Cells were incubated with 1.5 ml of a 1:1000 dilution of a mouse monoclonal IgG2a in PBS, pH 7.4 supplemented with 0.1% BSA for 30 min at 20°C. After three washes, cells were incubated with 1.5 ml of a 1:5000 dilution of goat anti-mouse IgG–F(ab')2–peroxidase conjugate (Immunotech, Marseilles, France) in PBS, pH 7.4 supplemented with 0.1% BSA for 30 min at 20°C. After three washes, cells were re-suspended in 0.1 M Tris–HCl, pH 8.5 and counted. For each sample, 2x105 cells per well were seeded in triplicate on 96-well plates (Nunc). Plates were developed with 150 µl of HPPA substrate at 26.5 mM (Sigma) diluted in 0.1 M Tris–HCl, pH 8.5, 0.13% Tween-20 supplemented with 3.75/10 000 (v/v) H2O2. After 30 min at 20°C, the reaction was stopped with 50 µl of 2 M glycine, pH 10.3. Fluorescence ({lambda}ex 330 nm, {lambda}em 425 nm ) was read in a Fluorolite 1000 plate fluorimeter (Dynatech Laboratories).

When applicable, modifications of the procedure are indicated in the Figure captions. For proteinase K treatment, A20 cells were incubated with 0.05 mg/ml of proteinase K (Sigma) in PBS, pH 7.4 at 37°C for 30 min. Cells were washed three times in PBS before incubation with the fusion proteins and vizualization was performed as described above.

Confocal microscopy

Cells L929 were plated on glass cover-slips (12 mm diameter in 24-well plates) the day before the experiment. They were washed twice with PBS and once with anchoring buffer (24.5 mM sodium citrate, 25.5 mM citric acid, pH 4.7, 280 mM sucrose) before incubation for 30 min at room temperature with T-ZZ or ZZ-T at 10–6 M in 0.4 ml of anchoring buffer. Control cells were treated with anchoring buffer only. For some experiments, cells were washed with PBS and cultivated for 24 h before transferrin incubation and processing for immunofluorescence. After two washes with PBS, cells were incubated with transferrin-FITC (25 µg/ml in DMEM containing 0.1 mg/ml BSA) for 45 min at 37°C. They were then cooled to 4°C, washed once with DMEM–BSA and twice with PBS. Cells were subsequently fixed with 3.7% paraformaldehyde (freshly prepared in PBS) for 30 min at room temperature. The fixative was quenched for 10 min with 50 mM ammonium chloride in PBS and the cover-slips were transferred to a humidified chamber and incubated for 45 min in permeabilization buffer (PBS supplemented with 0.02% saponin and 1 mg/ml BSA) containing rabbit anti-goat IgG (Nordic). After rinsing, the presence of the first antibody was revealed by incubation under the same conditions with a TRITC-labeled goat anti-rabbit IgG (Sigma). The cover-slips were then rinsed extensively with permeabilization buffer, then with PBS and briefly with water before mounting and examination under a Leica confocal microscope. Median optical sections were recorded with a x63 lens.

K562 cells growing in suspension were labeled with T-ZZ or ZZ-T and transferrin-FITC at a density of 5x106 cells/ml before processing for immunofluorescence (Subtil et al.1997Go) using the antibodies indicated above.

Cell permeabilization study

Vero cells were seeded on 96-well plates (Nunc) at a density of 105 cells/well in Dulbecco-MEM supplemented with 10% fetal calf serum (FCS) and 2 mM glutamine and were grown overnight to confluence. Cells were washed three times with medium without FCS. Then 200 µl of BCECF-AM solution (50 µg of lyophilized BCECF-AM (Molecular Probes) diluted with 72 µl of DMSO and 7.2 ml of medium without FCS) were added. Cells were then incubated for 1 h and washed three times with medium without FCS . A 50 µl volume of PBS, pH 7.4 was added to the cells and a first fluorescence measure ({lambda}ex 465 nm, {lambda}em 530 nm ) was done to define 100% fluorescence, using a Fluorolite 1000 plate fluorimeter. Medium was removed and recombinant proteins were added at the indicated concentrations in 50 µl of PBS adjusted to pH 4.8 with H2SO4. Fluorescence was monitored for 18 min. Then 0.5 µl of 1 M acetate, pH 3.6, which diffuses across cell membranes, was added (the final pH in the well remained at 4.8) to acidify the cytoplasm of cells. Fluorescence was further monitored for 5 min.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Recombinant proteins

We have produced three fusion proteins, in which the ZZ protein was fused to either the N- or C-terminus or to both extremities, of the T domain. These constructs are referred to as T-ZZ, ZZ-T and ZZ-T-ZZ. We have also produced the isolated T domain and ZZ protein for controls. Table IGo summarizes the primary structure of all five molecules. The 10 last amino acids of the T domain (from Arg377 to Thr386) are stretched out of its globular structure (Bennett and Eisenberg, 1994Go). In the T-ZZ protein, they constitute a spacer between the two fusion partners, together with two additional residues. In ZZ-T and ZZ-T-ZZ, a flexible hydrophilic peptide spacer rich in Gly and Ser residues was inserted between ZZ and T.

After expression and purification of the proteins, the final recovery yields ranged from 10 mg for T to 60 mg for ZZ-T-ZZ per liter of bacterial culture. SDS–PAGE and silver staining of the recombinant proteins showed that they had been purified nearly to homogeneity (Figure 1Go). They have an electrophoretic mobility consistent with the expected molecular masses deduced from their amino acid sequences: 16.74 kDa (149 residues) for ZZ, 22.01 kDa (201 residues) for T, 36.38 kDa (328 residues) for T-ZZ, 36.29 kDa (335 residues) for ZZ-T and 50.66 kDa (462 residues) for ZZ-T-ZZ. All proteins containing the ZZ part were capable of binding IgG. This was demonstrated by IgG affinity chromatography, the procedure used for their purification.



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Fig. 1. SDS–PAGE and silver staining of the recombinant proteins used in this study.

 
Interaction of the fusion proteins with the membranes of LUV

Interaction with membranes of the isolated T domain and related fusion proteins, as well as native diphtheria toxin, can be assessed in an assay measuring the release of a fluorescent dye entrapped into lipid vesicles (Zhan et al.1995Go; Liger et al.1998Go; Nizard et al.1998Go; Sharpe et al.1999Go). All proteins containing the T domain were able to permeabilize LUV at acidic pH (Figure 2Go) , whereas no permeabilization occurred at pH 7.4 (not shown). The isolated ZZ protein had no effect at either pH. Comparison of the effect of increasing concentrations of each protein shows that they all have comparable permeabilization efficiency on a molar basis and that this effect is dose dependent (Figure 2BGo). These results indicate that the capacity of the T domain to interact with membranes is not altered when another protein is fused to its N- or C-terminus or to both extremities.



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Fig. 2. Permeabilization of LUV by the recombinant proteins. (A) Comparison of the effect of each protein at 2 µM. (B) Comparison of the maximum effect reached after 140 s with various concentrations of T-ZZ (circles), ZZ-T (squares), ZZ-T-ZZ (triangles) and T (diamonds). 100% pyranine release was defined as the maximum of pyranine extinction observed with FCCP, a lipophilic ionophore for protons.

 
The capacity of these fusion proteins to interact with LUV and to bind IgG raises the possibility of using them to target LUV towards cells. This could be done by anchoring, via the T domain, to the surface of LUV IgG specific for cell surface markers. However, despite our efforts, targeting experiments gave inconsistent results. To investigate the reasons for this failure, we examined, using atomic force microscopy, the structure of the LUV after interaction with the fusion protein T-ZZ at acidic pH. Nice round-shaped LUV could be observed at acidic pH in the absence of protein (Figure 3AGo) and in the presence of the protein at pH 7.4 (not shown). However, in the presence of protein at acidic pH and at a lipid/protein ratio of 40 (w/w), massive fusion and fragmentation of LUV were observed (Figure 3B and CGo). These results indicate that the interaction of T domain fusion proteins destabilizes synthetic membranes. At a lipid/protein ratio of 400 (w/w), LUV kept their integrity with only small detectable changes in their shape (not shown). However, such a ratio was ineffective in promoting IgG-mediated binding of LUV to cells. Thus, membrane anchors for IgG derived from the T domain may not be used for the targeting of liposomes.



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Fig. 3. Structure of LUV incubated with the fusion protein T-ZZ at acidic pH observed by atomic force microscopy. (A) LUV in PBS–acetate at pH 4.8, without T-ZZ. Similar images were obtained for LUV incubated with PBS–acetate at pH 7.4, with or without T-ZZ. (B and C) LUV in PBS–acetate at pH 4.8, with T-ZZ at a lipid/protein ratio of 40 (w/w).

 
Binding of the fusion proteins to the surface of cells

We compared the binding capacities of the fusion proteins to the surface of L929 fibroblasts (Figure 4A and BGo) and A20 cells (Figure 4C and DGo). L929 cells attach to the plastic of the tissue culture plates, on to which they proliferate to confluence. A20 cells grow in suspension. The cells were incubated with varying concentrations of the fusion proteins at pH 4.8 for 1 h at 20°C. Binding to the cells was measured by immunoenzymatic detection using a mouse monoclonal IgG2a, capable of binding to ZZ by its Fc part and a goat anti-mouse IgG–F(ab')2–peroxidase conjugate. The results show that the fusion proteins T-ZZ and ZZ-T have the same capacity to bind to the surface of cells, in a dose-dependent manner. The same result was found for ZZ-T-ZZ with L929 cells. However, this triple fusion protein contains two ZZ moieties and should theoretically bind twice as much IgG as the two other fusion proteins. We found previously that all three proteins have the same capacity to interact with LUV. Thus, two possible explanations could account for this: either one of the ZZ moieties is masked by the other one within the bound ZZ-T-ZZ molecule, or once an IgG is bound to one ZZ it masks recognition of the second ZZ moiety by another IgG.



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Fig. 4. Binding of the fusion proteins T-ZZ, ZZ-T and ZZ-T-ZZ to the surface of cells studied by immunoenzymatic detection. In all experiments, cells were incubated with the proteins for 1 h at 20°C. After washing with PBS at pH 7.4, binding of the proteins was detected using a mouse monoclonal IgG2a and a goat anti-mouse IgG–peroxydase conjugate. Adherent fibroblastic L929 cells were incubated with various concentrations of the fusion proteins T-ZZ, ZZ-T and ZZ-T-ZZ at pH 4.8 (A) or 7.4 (B). As a control, cells were also incubated with a mixture of the isolated T domain and the ZZ protein, both at pH 4.8 and 7.4 (B). (C) Non-adherent A20 cells in suspension were incubated with various concentrations of the fusion proteins T-ZZ and ZZ-T at pH 4.8 or 7.4. (D) As a control, A20 cells were also incubated with a mixture of the isolated T domain and the ZZ protein, both at pH 4.8 and 7.4.

 
At pH 7.4, no binding to cells was observed with any of the fusion proteins. In addition, no binding of IgG to cells was found when cells were incubated with a mixture of the isolated T domain and the isolated ZZ protein, at neutral or acidic pH (Figure 4B and DGo).

Overall, these results show that the T domain may function as a membrane anchor for the ZZ protein fused at its N- or C-terminus. The capacity of these anchors to attach IgG to cells does not depend on the extremity of the T domain to which ZZ is fused. However, although T can be used to anchor two ZZ domains fused to both of its extremities, this does not lead to an increase in the amount of IgG anchored to cells. Optimal binding occurs for concentrations within the µM range using incubations of 1 h. This incubation time was chosen according to the kinetic study described in the next section.

Kinetics of binding to cells

We studied the influence of incubation time on the amount of fusion proteins bound to the surface of cells. L929 fibroblasts were incubated with the fusion proteins at pH 4.8 for various times before detection. The results in Figure 5AGo show that maximum binding is reached after 1 h. Half-maximum binding is reached between 10 and 20 min for T-ZZ and ZZ-T and after 30 min for ZZ-T-ZZ.



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Fig. 5. (A) Effect of incubation time on the binding of the fusion proteins T-ZZ, ZZ-T and ZZ-T-ZZ to the surface of L929 cells at pH 4.8. Cells were incubated for various times with the proteins at a concentration of 3x10–6 M at 20°C. After washing with PBS at pH 7.4, binding of the proteins was detected using a mouse monoclonal IgG2a and a goat anti-mouse IgG–peroxydase conjugate. (B) Life span of the fusion proteins T-ZZ, ZZ-T and ZZ-T-ZZ once bound to the surface of L929 cells. The proteins were incubated with the cells for 1 h at a concentration of 3x10–6 M at 20°C, at pH 4.8. After washing with PBS at pH 7.4, the cells were incubated with culture medium in the CO2 incubator at 37°C. After various times, bound proteins were detected with an IgG2a and then with an anti-IgG–peroxydase conjugate.

 
Fate of the fusion proteins bound to the cell surface

Our aim was to investigate the fate of the fusion proteins bound to the surface of live cells. L929 fibroblasts were treated with the fusion proteins at acidic pH for 1 h at 20°C, to allow binding of the proteins to the surface of the cells. The cells were then washed and placed back into culture medium containing FCS, at 37°C, in the CO2 incubator. The presence of the fusion proteins remaining at the surface of the cells was revealed after various times. Figure 5BGo shows that about 75% of the proteins were still displayed at the cell surface after 24 h.

Confocal microscopy was then used to study the subcellular localization of the fusion proteins. L929 fibroblasts were incubated with T-ZZ or ZZ-T at acidic pH for 30 min, washed and then incubated with FITC-labeled transferrin for 45 min. For confocal observations, the cells were fixed, permeabilized and the fusion proteins were detected using a rabbit anti-goat IgG and a TRITC-labeled goat anti-rabbit IgG. Transferrin is known to be rapidly internalized through the clathrin-coated pathway and to accumulate in early endosomes (Subtil et al.1997Go). Transferrin–FITC was therefore used as an endosome marker. Figure 6A–C Go shows that transferrin (green) is internalized and localized to the perinuclear area, as expected (Subtil et al.1997Go). Identical images were obtained when the transferrin receptor was revealed by immunofluorescence on permeabilized cells (not shown), indicating that the permeabilization procedure was adequate. Nevertheless, the T-ZZ and ZZ-T molecules (red) were observed at the cell surface and were not internalized.



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Fig. 6. Confocal microscopic analysis of cells treated with the fusion proteins T-ZZ and ZZ-T at acidic pH. L929 cells (AE) and K562 cells (FH) were incubated with T-ZZ (B, D, G), ZZ-T (C, E, H) or no protein (A, F) at pH 4.8 and room temperature for 30 min and then washed thoroughly. Cells were then incubated with FITC-labeled transferrin (green) for 45 min at 37°C. After fixation and permeabilization of the cells, the fusion proteins were detected using rabbit IgG and a TRITC-labeled goat antibody anti-rabbit IgG (red). In (D) and (E), L929 cells were cultured for 24 h before incubation with transferrin and detection of the fusion proteins. Median optical sections were obtained using a Leica confocal microscope. Bar = 5 µm.

 
To examine whether some internalization could take place with time, cells were loaded with the fusion proteins and cultured for 24 h, before labeling with transferrin–FITC for 45 min and processing for immunofluorescence. The results show (Figure 6D and EGo) that the proteins are not internalized after 24 h and that an important proportion of the molecules is still present at the surface of the cells. As expected, transferrin-FITC was found in intracellular compartments, showing that the cells were still alive and retained their ability to internalize.

Similar images were obtained when a T domain–cytokine fusion protein was anchored on to RMA lymphoma cells, B16 melanoma cells, and L929 cells (unpublished results). Thus, it seems that persistence of T domain fusion proteins at the surface of live cells is a general phenomenon. However, the study of the binding of the fusion proteins T-ZZ and ZZ-T on the K562 human myeloid leukemia cell line gave different results. In this case, the fusion proteins were internalized and gathered into the same compartments as transferrin–FITC, as shown by the yellow color resulting from green and red dye co-localization (Figure 6G and HGo). This compartment is presumably the early endosome. A fraction of the proteins remained at the surface of the cells after 45 min (Figure 6G and HGo). Thus, internalization of the proteins seems slower than receptor-mediated internalization of transferrin. No protein was detectable either at the surface or into cell compartments after 24 h (not shown).

Overall, our results show that the fate of T fusion proteins bound to the surface of cells may depend on cell type. On cells that do not internalize the T fusion proteins, they remain displayed at the cell surface for more than 24 h, without much loss. On cells that do internalize the fusion proteins, they disappear from the surface more slowly than transferrin bound to its receptor and seem to gather in the early endosomes in the same manner as transferrin. After 24 h, no protein remains detectable on these cells.

Cell surface proteins are not required for binding of the fusion proteins to the cell membrane

So far, we have found that the fusion proteins described in this work, as well as T-cytokine fusion proteins, could bind to any cell type. Up to nine different cell types have been tried, giving similar results. These include fibroblasts, leukemia cell lines from various lineages, breast cancer and melanoma cell lines. This makes sense if one considers that the T domain has affinity for phospholipid bilayers. However, we investigated the possible implication of cell surface proteins in the binding of T-fusion proteins to cells. Cells were treated with proteinase K to remove extracellular domains of membrane proteins. The data in Figure 7AGo show that T-ZZ at acidic pH has the same capacity to bind to cells whether treated or not with proteinase K. This strongly suggests that the extracellular domains of cell surface proteins do not influence binding of T to the lipid bilayer of the membrane.



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Fig. 7. (A) Binding of the fusion protein T-ZZ to the surface of A20 cells treated with proteinase K. Cells treated or not with proteinase K were incubated with various concentrations of T-ZZ. Binding of the protein was detected by incubation using a mouse monoclonal IgG2a and a goat anti-mouse IgG–peroxydase conjugate. (B) Analysis of membrane permeability to H+ of Vero cells treated with the fusion proteins at pH 4.8. Cells were loaded with the fluorescent intracellular pH probe BCECF-AM. Fluorescence intensity of the cells in PBS at pH 7.4 defined maximum fluorescence (100%). Cells were then incubated in PBS at pH 4.8 with the fusion proteins at a concentration of 0.8x10–6 M or with the isolated T domain at a concentration of 1.3x10–6 M. In controls, cells were incubated with PBS at pH 4.8 with the isolated ZZ protein at a concentration of 1.3x10–6 M or with no protein. An additional control was made by incubating cells with acetate buffer at pH 4.8, which rapidly penetrates cells and acidifies the cytoplasm. Fluorescence intensities were monitored for 23 min and normalized. After 18 min, acetate buffer was added to all cells to acidify their cytoplasm.

 
Binding of the fusion proteins to cells does not modify membrane permeability

We have shown previously that the interaction of the T domain and its related fusion proteins, with LUV at acidic pH modifies their permeability dramatically (Figure 2Go). It was therefore important to investigate whether the T domain fusion proteins alter the permeability of cell membranes. Indeed, it has been shown that interaction of the complete diphtheria toxin with the membrane of Vero cells at acidic pH could lead to the formation of ion channels under certain conditions (Papini et al.1988Go; Sandvig and Olsnes, 1988Go; Stenmark et al.1988Go). These channels were found to be selective for small cations such as H+, Na+ and K+. To examine this question, Vero cells were loaded with BCECF, a fluorescent intracellular pH probe, using the acetoxymethyl ester loading method. The cells were incubated with the fusion proteins at pH 4.8 and their fluorescence was monitored. In case of increased membrane permeability, H+ ions would enter the cells and acidify their cytoplasm, leading to a decrease in fluorescence. In the absence of protein, a loss of fluorescence intensity of 25% was observed, indicating a moderate acidification of the cytoplasm of the cells (Figure 7BGo). No further acidification of the cytoplasm was observed in the presence of any of the T fusion proteins or the isolated T domain over 12 min. As a positive control for cytoplasm acidification, an acetate buffer at pH 4.8 was used. Acetate, diffusing through the cell membrane, resulted in a 50% decrease in fluorescence intensity within 3 min. Overall, these results strongly suggest that binding of the T domain fusion proteins, as well as the isolated T domain, at acidic pH, does not increase cell membrane permeability to H+. In addition, no alteration of cell shape or cell membrane bursting was ever observed by microscopy following this treatment.


    Discussion
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
This work shows that the T domain may be used as a membrane anchor for soluble proteins fused at its N- or C-terminus. The capacity of interaction of the T domain with membranes does not depend on which of its extremities the soluble protein is linked (Figure 2Go). This allows one to choose the orientation of the protein to be attached to membranes. For instance, the recognition site of that protein could be orientated facing away from the membrane, to allow interaction with a receptor carried by another cell. The T domain may also be used to anchor two soluble proteins at a time, one at each of its extremities (Figure 4AGo). However, in this work, recognition of the triple fusion protein ZZ-T-ZZ by IgG at the cell surface was not stronger than for the two other fusion proteins carrying only one ZZ protein. Several explanations could account for this, considering the spatial proximity of both ZZ moieties within ZZ-T-ZZ. One ZZ could mask the other one. Once an IgG is bound to one ZZ it could mask recognition of the second ZZ by another IgG. Finally, one IgG could bind both ZZ, as the Fc region carries two ZZ-binding sites (Deisenhofer, 1981Go). Further investigation will be needed to establish the potential of linking two different proteins to the T domain.

The N-terminal side of T plays an important role in the translocation process of the C domain within the native toxin (Madshus, 1994Go; Madshus et al.1994Go). However, no membrane insertion or translocation of ZZ seems to occur when anchoring the fusion protein ZZ-T to cells. Indeed, its ZZ part is recognized by IgG in a similar fashion as for T-ZZ. It has been shown previously that binding of the toxin to its receptor was a prerequisite for C domain entry into cells from the cell surface at acidic pH (Stenmark et al.1988Go; Madshus et al.1991Go). In our case, no protein receptor is involved in the interaction with the membrane. In addition, C must change its conformation in order to interact with T, insert into the membrane and translocate (Falnes and Olsnes, 1995Go; Ren et al.1999Go). ZZ, being very stable and soluble (Samuelsson et al.1994Go), is not suited to membrane interaction or translocation.

A striking result is that whereas lipid vesicles are destabilized by the interaction of the membrane anchors (Figure 3Go), the cell membrane seems not to be affected. Cells remain fully viable (Nizard et al.1998, and unpublished results ), their membranes are not permeabilized (Figure 7Go), no alteration of cell morphology was observed upon confocal microscopic examination (Figure 6Go) and in atomic force microscopy studies of cell surface (not shown). This is perhaps not surprising. The cell membrane is far more complex than the membrane of LUV. It has a very different lipid composition including cholesterol, leaflet asymmetry and transmembrane domains of proteins and is connected to a cytoskeleton. The fusion of lipid vesicles induced by diphtheria toxin at acidic pH has already been described using fluorescence resonance energy transfer (Papini et al.1987Go). Here, we illustrated this phenomenon by showing the extent of LUV structure alteration (Figure 3Go). This result may be interesting for those studying the effect of hemolytic toxins on lipid vesicles, as the nature of the lesions in the membranes of cells or vesicles may be very different.

Finally, we found that the fate of the fusion proteins anchored to the surface of cells may differ, depending on the cell type. On fibroblasts (this study), lymphoma cells and melanoma cells (unpublished results), the proteins remain associated with the surface of the cells for at least 24 h without much loss. In sharp contrast, K562 cells internalize the proteins, almost as efficiently as transferrin. So far, we have studied binding of T-derived fusion proteins to the surface of nine different cell lines and have not found important differences in binding levels. Although it is necessary to check, for any new cell type, the behavior of the fusion proteins, it is possible that most cells will not internalize them. This difference in behavior may lead to different applications for the use of T domain membrane anchors. In the case where fusion proteins remain at the plasma membrane, it is possible to display a protein at the surface of cells for maybe several days. This protein could act as a signal for or promote interaction with other cells. It could also be used as a marker to follow a labeled cell in a population. The pathway through which some cell types internalize the fusion proteins could be used to deliver such proteins to the endosomes. However, this pathway will need further characterization.

Other membrane anchors have been described to attach proteins to cells. Chelator lipids can be incorporated into the cell membrane to attach histidine-tagged proteins using nickel ions (van Broekhoven et al.2000Go). A glycosylphosphatidylinositol anchor (McHugh et al.1999Go) and an acylated tag anchor (de Kruif et al.2000Go) have also been described. Each system may have its own advantages depending on the application, the ease of tagged protein expression and purification, the conditions of anchoring needed (dead or live cells, need for a triggered anchoring such as the pH for T), the persistence on the cell surface, etc.


    Notes
 
4 To whom correspondence should be addressed. E-mail: daniel.gillet{at}cea.fr Back


    Acknowledgments
 
We thank Dr Florent Guillain for help in setting the atomic force microscopy study and Dr Bernard Maillere for critical comments and discussion. P.N. and A.C. were supported by the Ministère de l'Education Nationale, de la Recherche et de la Technologie. This work was supported by the Comité Départemental de l'Essonne de la Ligue Nationale Contre le Cancer, the Association pour la Recherche contre le Cancer and the Commissariat à l'Energie Atomique.


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Received November 20, 2000; revised March 3, 2001; accepted March 20, 2001.