Yuwen Meng
email: yuwen.meng.etu@univ-lemans.fr

Erwan Nicol
email: erwan.nicol@univ-lemans.fr

Taco Nicolai
email: taco.nicolai@univ-lemans.fr

Exploiting multiple phase separation to stabilize water in water emulsions and form stable microcapsules

Keywords: Aqueous multiphase system, microcapsule, gelatin, emulsion

Hypothesis

Water in water (W/W) emulsions are formed by mixing aqueous solutions of incompatible polymers.

It is possible to add a third polymer solution that forms at the right conditions a phase that completely covers the dispersed droplets as a thin layer. Our hypothesis is that by gelling the third phase, W/W emulsions can be stabilized and that microcapsules can be formed that are stable against dilution. Experiments W/W emulsions were formed by mixing aqueous solutions of poly(ethylene oxide) (PEO) and dextran.

Gelatin was added to form the third phase, gelation of which was induced by cooling. The morphology was observed by microscopy, and the rheological properties were investigated.

Findings

The compatibility of gelatin and PEO can be fine-tuned by the pH such that a continuous layer of the gelatin phase forms around the droplets of the dextran phase, with a thickness that can be varied.

After cooling, the gelatin layer forms a gel and provides stabilization against coalescence. The gelatin microcapsule was found to be stable to dilution. The generality of the method was demonstrated by applying it to another, fully food-grade, W/W emulsion formed by mixing amylopectin and xyloglucan.

Introduction

When two aqueous solutions of incompatible polymers are mixed, a water in water (W/W) emulsion is formed with the minority phase dispersed in the form of droplets in a continuous medium of the majority phase [START_REF] Frith | Mixed biopolymer aqueous solutions -phase behaviour and rheology[END_REF]. When more than two types of incompatible polymers are mixed, droplets of different phases are dispersed in the continuous phase. In this manner water in water in water (W/W/W) emulsions can be formed. W/W/W emulsions have been little investigated so far.

Song et al. [START_REF] Song | Monodisperse w/w/w double emulsion induced by phase separation[END_REF][START_REF] Song | All-aqueous multiphase microfluidics[END_REF] used microfluidics to form an A/B/A emulsion consisting of droplets of a phase rich in dextran (DEX) dispersed in a continuous phase rich in poly(ethylene oxide) (PEO) with smaller droplets of the PEO phase within the DEX droplets. A different method, not involving microfluidics, was used by Beldegrun et al. [START_REF] Beldengrün | Formation and stabilization of multiple water-in-water-in-water (W/W/W) emulsions[END_REF] to produce A/B/A emulsions consisting of droplets of a phase rich in gelatin (GEL) dispersed in a continuous phase rich in maltodextrin with smaller droplets of the maltodextrin phase within the GEL droplets.

Very recently, Nicolai et al. [START_REF] Rowlinson | Molecular theory of capillarity[END_REF] reported an investigation of A/B/C emulsions prepared by simply mixing three different incompatible macromolecules forming the three phases. They showed that droplets of phases A and B in a continuous phase C may spontaneously stick to each other. This happens if none of the interfacial tensions of the three interfaces ( AB ,  AC ,  BC ) is larger than the sum of the interfacial tensions of the other two interfaces [START_REF] Rowlinson | Molecular theory of capillarity[END_REF]. When one interfacial tension is larger than the sum of the other two, the two phases with the largest interfacial tension will be separated by the third phase. The angle () that each phase makes with the other two phases depends on the relative interfacial tensions:

If the interfacial tension between two phases is known, that between the other phases can be determined by measuring the angles. In this manner, Nicolai et al. [START_REF] Nicolai | Effect of the Interfacial Tension on Droplet Association in Aqueous Multiphase Systems[END_REF] determined the interfacial tension between 14 different pairs of aqueous polymer phases with respect to that between DEX and PEO.

Obviously, droplets of the same phase coalesce as soon as they collide. Therefore, aqueous multiphase systems are unstable and eventually, macroscopic phase separation occurs. Molecular surfactants cannot be used to stabilize W/W emulsions, because they cannot effectively span the width of the W/W interface, which is determined by the correlation length of the polymer solutions, i.e., several nm. In addition, the interfacial tension of W/W interfaces, which drives the adsorption, is much lower than that of the oil/water interface [START_REF] Hans Tromp | Composition, concentration and charge profiles of water-water interfaces[END_REF][START_REF] Scholten | Interfacial tension of a decomposed biopolymer mixture[END_REF]. However, it has been shown that W/W emulsions can be stabilized by adding particles [START_REF] Nicolai | Particle stabilized water in water emulsions[END_REF][START_REF] Esquena | Water-in-water (W/W) emulsions[END_REF][START_REF] Dickinson | Particle-based stabilization of water-in-water emulsions containing mixed biopolymers[END_REF] or some types of linear polyelectrolytes [START_REF] Tea | Stabilization of Water-In-Water Emulsions by Linear Homo-Polyelectrolytes[END_REF][START_REF] Meng | Utilization of xanthan to stabilize water in water emulsions and modulate their viscosity[END_REF] that spontaneously adsorb at the interface forming a layer that inhibits coalescence. It was shown that aqueous multiphase systems can also be stabilized in this manner [START_REF] Nicolai | Effect of the Interfacial Tension on Droplet Association in Aqueous Multiphase Systems[END_REF]. The difficulty with this method is, however, that it is still unclear what properties the particles should have to stabilize a particular W/W emulsion, as it was found that it is not enough that the particles adsorb at the interface. It appears that a certain amount of interaction between the particles at the interface is also required [START_REF] Machado | Effect of adding a third polysaccharide on the adsorption of protein microgels at the interface of polysaccharide-based water in water emulsions[END_REF][START_REF] Machado | Stabilization of amylopectinpullulan water in water emulsions by Interacting protein particles[END_REF]. Furthermore, stabilization becomes very difficult with dilution closer to the binodal where the interfacial tension goes to zero.

Here we propose a different method to stabilize W/W emulsions by adding a small amount of a third phase B. If the conditions are such that  AC >  AB + BC , phase B surrounds the dispersed droplets of phase A, separating them from the continuous phase (C). Coalescence of droplets can be inhibited by subsequently gelling the layer of phase B. Of course, once the layer has gelled, there is no longer a need for the interfacial tension between the phases, and the layer will persist even after dilution below the binodal. Therefore, this method provides the means to form stable microcapsules of phase B within a fully aqueous system. This cannot be done with W/W emulsions stabilized by particles as the particles simply redisperse below the binodal. The method to form stable microcapsules reported in the literature involved using oil in water emulsions [START_REF] Lazko | Soy glycinin microcapsules by simple coacervation method[END_REF][START_REF] Jaganathan | Protein microcapsules: preparation and applications[END_REF]. The advantage of using W/W emulsions is that the oil phase does not need to be removed, which may be difficult. In addition, water-soluble ingredients can be easily encapsulated using W/W emulsions.

We have investigated the feasibility of our new approach using a model W/W emulsion formed by mixing DEX and PEO in which the DEX phase is dispersed as droplets. GEL was added to form the third phase, because GEL solutions are liquid at temperatures above 30 °C and quickly jellify when cooled to 10 °C. The generality of the method was tested by investigating the fully food-grade W/W emulsion formed by mixing the polysaccharides amylopectin and xyloglucan in the presence of gelatin.

Materials and Methods

Materials

PEO (M w = 2.0 × 10 5 g.mol -1 ), dextran (M w = 1.6 × 10 5 g.mol -1 ), gelatin from bovine skins (Type B, 225 g Bloom), gelatin from cold water fish skin (M w = 4.4 × 10 5 g.mol -1 ), amylopectin (AMP) from maize (M w = 1.0 × 10 8 g.mol -1 ), the fluorescent dye rhodamine B and dextran labelled with fluorescein isothiocyanate (FITC-DEX) were all purchased from Sigma-Aldrich. Xyloglucan (XYL) with M w = 1.0 × 10 6 g.mol -1 was kindly provided by DSP Gokyo Food & Chemical co. The amylopectin was purified according to a protocol provided in the literature [START_REF] Bello-Pérez | Macromolecular Features of Starches Determined by Aqueous High-performance Size Exclusion Chromatography[END_REF]. Stock solutions of Dex, PEO, Gel, AMP and XYL were prepared with ultrapure water (MilliQ) at 30, 15, 20, 15 and 2 wt%, respectively. The PEO powder contained a very small weight fraction of silica particles that were removed by centrifugation at 5 × 10 4 g for 4.5 h. GEL was solubilized at 80 °C. The XYL solution was centrifuged at 5 × 10 4 g for 30 min to remove insoluble particulates. The pH of the solutions was measured using a pH meter and adjusted by dropwise addition of HCl or NaOH at 0.1 or 1 mol.L -1 .

Methods

Emulsions were prepared by mixing the stock solutions in the required amounts at 40 °C using a vortex stirrer. The morphology and behavior of the emulsions did not depend on the order of mixing nor on the duration and speed of stirring.

The concentration of GEL in each phase was determined after macroscopic phase separation by measuring the absorption of UV light at a wavelength of 275 nm using a UV-light spectrometer (Jasco PAC-743). The extinction coefficient of GEL was determined by measuring a range of known concentrations and was found to be 0.102 g -1 •L•cm -1 .

Confocal laser scanning microscopy (CLSM) measurements were done at 20 °C with a Zeiss LSM800 CLSM microscope (Carl Zeiss Microscopy GmbH, Germany) utilizing two different water immersion objectives: HC × PL APO 63× (NA = 1.2) and a HC × PL APO 25× (NA = 0.7). The samples were loaded into acrylic wells microplates and labeled by adding 5 ppm of rhodamine B, which was excited at 561 nm. The rhodamine B partitioned strongly to the GEL phase in mixtures with PEO and DEX with the excess partitioning preferentially to the PEO phase. In a few cases, a small amount of FITC-DEX was added in order to measure the release of DEX after dilution. ImageJ software was used to process the images. Shear measurements were done using a stress-controlled rheometer (ARG2, TA instruments) with a cone and plate geometry (radius 40 mm, angle 2°, gap 56 mm). Gelation of aqueous GEL solutions was monitored during cooling and subsequent heating ramps between 10 and 60 °C at 0.5 °C.min -1 at an oscillation frequency of f = 1 Hz. The applied deformation (10 % or 1 %) was chosen within the linear response regime. The gelation kinetics were studied by measuring the storage (G' ) and loss (G") moduli as a function of time at 10 °C. Gelation of the emulsions was investigated at 10 °C and 20 °C at f = 0.1 Hz.

Results and discussion

Morphology of emulsions in the presence of fish gelatin

Gelatin from cold water fish skin does not form a gel in aqueous solution at room temperature, making it easier to study the morphology of the emulsions with liquid PEO, DEX and Gel phases. Mixtures of 10 wt% PEO, 2.6 wt% DEX and 4 wt% GEL phase separated into PEO, DEX and Gel rich phases with volume fractions of 80 %, 10 %, and 10 %, respectively. The angles () that each phase made with the two other phases were determined in the same manner as explained elsewhere [START_REF] Nicolai | Effect of the Interfacial Tension on Droplet Association in Aqueous Multiphase Systems[END_REF] and are plotted as a function of the pH in Fig. 2. They were approximately constant between pH 5 and pH 9, but  increased sharply to 180° for the PEO and DEX phases at low and high pH and decreased sharply to 0° for the GEL phase. This pH dependence can be understood by the fact that the GEL and PEO phases started to mix at low and high pH, which strongly reduced the interfacial tension between GEL and PEO ( GP ). In a narrow pH range,  GP was sufficiently small so that  DP >  GD +  GP , which is the condition that is required to form a continuous layer of the GEL phase around the droplets of the DEX phase. The relationship between the charge density of GEL and the pH was determined by titrating a neat GEL solution with NaOH or HCl. A strong pH dependence on the amount of added HCl or NaOH was observed between pH 5 and pH 9 with a weaker dependence at lower and higher pH. The inflection point was situated at pH  7, which we consider to be the pH at which the net charge of the GEL is close to zero. In this manner, the number of charges per gram gelatin at each pH could be calculated from the amount of HCl or NaOH per gram gelatin that was needed to reach this pH neglecting the tiny amount needed to change the pH itself. Fig. 3 shows the number of charged groups per gram GEL as a function of the pH, which was found to be the same for fish GEL and bovine GEL within the experimental error. The effect of the pH on the phase volumes was investigated by centrifuging at 30 g until complete phase separation. The relative phase volume of the DEX phase did not depend significantly on the pH, but the GEL phase volume decreased below pH 6 and above pH 9, see Fig. S1 of the supplementary information. This observation shows that GEL dissolved into the PEO continuous phase at high and low pH, with only a small quantity going into the DEX phase.

Stabilization by gelation of the gelatin layer

Morphology of emulsions with bovine gelatin

Emulsions with bovine gelatin were prepared in the same manner as with fish gelatin, but at 40 °C to keep the GEL phase liquid. CLSM images were obtained immediately after mixing while the temperature rapidly decreased to 20 °C. The effect of the pH on the morphology of the emulsions with bovine GEL was similar to that observed with fish GEL discussed above, see Fig. S2 of the supplementary information. However, continuous GEL layers were formed with bovine GEL close to pH 3.8 and pH 10 compared with pH 4 and pH 11 for fish GEL. The difference may be due to differences in the composition and molar mass of the two types of GEL. When the emulsion at pH 10 was diluted, the GEL dissolved in the PEO phase. However, a continuous GEL layer was formed again when the pH was slightly reduced. This is illustrated in Fig. 4 which shows CLSM images of emulsions that were diluted by a factor (D) 1.5, 2 and 3. After dilution at D = 3, the volume fractions of both the DEX and GEL phases became small and it was impossible to form a continuous layer of GEL around the DEX droplets. The thickness of the GEL layer could be varied by varying the amount of GEL while keeping the concentrations of PEO and DEX the same, see Fig. 5. As might be expected, the thickness of the GEL layer increased with increasing GEL concentration. However, large GEL droplets containing several DEX domains were observed at higher GEL concentrations.

Stabilization by gelation of the bovine GEL layer

It is well known that aqueous solutions of bovine GEL form gels when cooled and become liquid again when heated [START_REF] Djabourov | Gelation of aqueous gelatin solutions. I. Structural investigation[END_REF]. This is illustrated in Fig. S3a of the supplementary information that shows the shear moduli of an aqueous GEL solution at 15 wt% as a function of the temperature during cooling and heating at pH 7. Similar results were obtained at other pH values. The gel and melt temperatures were defined as the temperature at which G' and G" crossed during cooling and subsequent heating. They were found to be approximately 22 °C and 30 °C, respectively, with very little dependence on the pH in the range 3 -10, see in Fig. S3b of the supplementary information. The gelation rate at 10 °C was studied by measuring the moduli as a function of time after rapidly reducing the temperature from 50 °C to 10 °C, see S4a of the supplementary Information. Gels were formed very rapidly, and subsequently, G' continued to increase slowly. G'  10 4 Pa after 10 min at 10 °C without a systematic dependence on the pH, see fig. S4b of the supplementary information.

The concentration dependence of G' measured after 10 min at 10 °C is shown in Fig. 6. G' increases sharply starting at C  1 wt% and increases more slowly for C > 5 wt%. Trials showed that gels were also formed at temperatures up to 25 °C, but the kinetics became increasingly slow with increasing temperature.

Fig. 6 Concentration dependence of the storage modulus of gels formed by bovine GEL solutions after 10 min at 10 °C. The solid line is a guide to the eye.

It is clear that the GEL layer covering the DEX droplets rapidly jellifies after cooling to 10°C.

Emulsions that had been cooled for 1 h in a refrigerator at 8 -9 °C were found to be stable for at least one week at 20 °C except the one at D = 3, in which case large DEX domains were formed together with aggregates of deformed GEL layers, see Fig. 4.

Dynamic mechanical properties

Not all GEL was situated in the layer as a significant fraction of the GEL remained dissolved in the PEO phase (Fig. S1). The concentration of GEL in the PEO phase was determined by measuring the UV light absorption after macroscopic phase separation. It was found to be 2.2, 1.6 and 1.2 wt% for D = 1, 1.5 and 2, respectively, which corresponded to about 55% of the GEL in the system. The presence of this relatively large fraction of the GEL in the PEO phase can be explained by the molar mass dispersity of the GEL with the smaller GEL chains remaining solubilized in the PEO phase. The residual GEL in the continuous PEO phase caused the emulsions to form weak gels at 10 °C, see Fig. S5 of the supplementary information, with the dispersed droplets trapped in the GEL network, which inhibited their sedimentation.

The weak GEL network could be easily broken by applying a shear force resulting in a viscous liquid. Fig. 7 shows the viscosity as a function of the shear rate of the emulsions at 10 °C. first increase of the shear rate, the GEL network is broken, which explains the maximum of the apparent viscosity at low shear rates. Repeated shear ramps did not show this maximum, because the network does not recover rapidly at 20 °C. As might be expected, the dynamic viscosity decreased when the system was diluted so that the continuous phase contained less GEL and PEO. A more detailed investigation of the shear rate dependence of the viscosity of W/W emulsions is beyond the scope of the present paper, but it has been addressed elsewhere [START_REF] Tea | Viscosity and Morphology of Water-in-Water Emulsions: The Effect of Different Biopolymer Stabilizers[END_REF]. The GEL layer was found to be quite resistant to shear, as the droplets were hardly deformed and remained well covered with the GEL layer. After stirring the emulsions with a magnetic stirrer at rates up to 1000 rmp the GEL layer resisted showing only small deformations at the highest stirring rates.

Dilution after cooling

Emulsions prepared at dilution factor D = 1.5, i.e., containing 7.2 wt% PEO, 1.3 wt% DEX and 2.6 wt% GEL, were cooled in the refrigerator for 1 h. They were subsequently diluted at 20 °C with MilliQ water and homogenized by stirring at 200 rmp during 30 s. The effect of dilution on the morphology is shown in Fig. 8. The difference between preparing the emulsions at different dilution factors and diluting after cooling is striking. Emulsions prepared at D = 3 did not form a continuous GEL layer (see Fig. 4), but the GEL layer resisted when the dilution was done after cooling. In fact, the GEL layer resisted even after dilution by a factor 6, i.e., when DEX and PEO no longer phase separated. No coalescence was observed, but in a few instances droplets associated without the GEL layer being broken. The droplets appeared to swell after dilution, and the GEL layer became thicker, which we attribute to the decreased osmotic pressure of the PEO phase after dilution. The diluted emulsions were stable for a period of at least one week, but slow sedimentation of droplets occurred, indicating that the residual GEL in the continuous phase was no longer capable of forming a network that could trap the droplets. As was mentioned above, DEX is miscible with PEO at D = 6 [START_REF] Gonzalez-Jordan | Influence of the Protein Particle Morphology and Partitioning on the Behavior of Particle-Stabilized Water-in-Water Emulsions[END_REF]. In order to check whether the DEX remained trapped in the droplets, a small amount of fluorescently labeled FITC-DEX was added with the same molar mass. Fig. 8 shows that DEX could escape through the GEL layer and become homogeneously dispersed after one day. More detailed observations as a function of time showed that complete release of DEX took several hours, see Fig. S6 of the supplementary information.

D = 2 D = 3 D = 6

Discussion

The method to stabilize W/W emulsions and form stable capsules was demonstrated here for a particular system for which we exploited the fact that the GEL phase forms a gel upon cooling.

Emulsions with a continuous GEL layer surrounding the dispersed droplets could only be formed close to pH 4 and pH 10. However, once the emulsions formed at pH 10 were stabilized by cooling, the pH could be lowered by adding dropwise a concentrated solution of HCl while stirring. Emulsions at pH 7 were also found to be stable during at least one week, and the GEL microcapsules were also stable after dilution, see Fig. S7 of the supplementary information.

In principle, any W/W emulsion can be stabilized by GEL in this manner as long as the conditions mentioned in the Introduction are fulfilled, i.e., the interfacial tension between the dispersed phase and the continuous phase needs to be larger than the sum of the interfacial tensions with the GEL phase. To test the universality of the method, we have investigated a fully food-grade W/W emulsion formed by mixing amylopectin (AMP) and xyloglucan (XYL). Emulsions prepared by mixing 6% wt% AMP, 0.48% wt% XYL and 1.75% wt% GEL that phase separated into AMP, XYL and GEL rich phases with volume fractions of 68 %, 14 %, and 18 %, respectively. In this case, dispersed droplets of the XYL phase were covered with a layer of the GEL phase at pH  7.5. After cooling, the emulsion was stable for at least a week at 20°C and resisted dilution below the binodal of AMP and XYL, see Fig. 9. Notice that small domains of the GEL phase were also formed during cooling within the AMP and XYL phases for this system. A further example of a food-grade W/W emulsion is formed by mixing guar and amylopectin [START_REF] Chen | Water-in-water-in-water emulsions formed by cooling mixtures of guar, amylopectin and gelatin[END_REF]. However, in this case, the formation of the Gel layer was not induced by varying the pH, but by cooling at a fixed pH, inducing concomitantly phase formation of the GEL phase and its gelation.

As was mentioned in the Introduction, it has been reported that stable protein microcapsules can be formed in a similar manner using oil in water emulsions [START_REF] Lazko | Soy glycinin microcapsules by simple coacervation method[END_REF][START_REF] Jaganathan | Protein microcapsules: preparation and applications[END_REF]. In this case, the proteins form a layer at the oil/water interface that is subsequently cross-linked to form a gel. The advantage of using W/W emulsions is that the oil phase does not need to be removed, which may be difficult. In addition, water-soluble ingredients can be easily encapsulated using W/W emulsions.

Conclusion

Building on previous work [START_REF] Machado | Effect of adding a third polysaccharide on the adsorption of protein microgels at the interface of polysaccharide-based water in water emulsions[END_REF], we have shown that the association between dispersed dextran and gelatin phases in a continuous PEO phase is determined by the relative interfacial tensions between the phases and can be varied by varying the pH. Importantly, by fine-tuning the pH close to pH 4 or pH 10, a continuous gelatin layer can be formed completely covering the dextran droplets. The gelatin layer can be jellified by cooling below 25°C, making the emulsions stable against agitation and dilution. Excess gelatin in the continuous phase can form a very weak gel that inhibits sedimentation of the droplets, but the emulsion still flows easily when weak stresses are applied. The pH of the cooled emulsion can be varied by adding acid or base while stirring. The gelatin microcapsules are stable after dilution even when PEO and dextran are no longer incompatible, in which case it was found that dextran was released into the continuous medium over a period of hours.

As hypothesized, multiphase separation provides a general method to stabilize W/W emulsions by adding a third phase that covers the dispersed droplets and that can be jellified. This new method provides a versatile alternative to adding particles, which has been done so far [START_REF] Nicolai | Particle stabilized water in water emulsions[END_REF][START_REF] Esquena | Water-in-water (W/W) emulsions[END_REF][START_REF] Dickinson | Particle-based stabilization of water-in-water emulsions containing mixed biopolymers[END_REF],

and provides a better stability of W/W emulsions in particular close to the binodal. It can be applied to fully food-grade systems as we showed here for amylopectin/xyloglucan mixtures. In agreement with our second hypothesis, the method is also an alternative route to form microcapsules in fully aqueous systems without the need to remove an oil phase as is required for the methods proposed in the literature [START_REF] Lazko | Soy glycinin microcapsules by simple coacervation method[END_REF][START_REF] Jaganathan | Protein microcapsules: preparation and applications[END_REF]. GEL microcapsules dissolve when heated above 30 °C, which may be useful for certain applications. However, stability at higher temperatures can potentially be obtained by enzymatic crosslinking the GEL layer using transglutaminase. The potential of the microcapsules formed in this manner for encapsulation and controlled release will need to be studied in future work. Extension to other gel forming aqueous phases will also need further investigation.

Fig. 1

 1 shows CLSM images of the emulsions at different pH immediately after preparation. At extreme low and high pH values (3 and 11.7), only two phases could be distinguished as GEL was dissolved into the continuous PEO phase. At intermediate pH (4.5 -10.8), droplets of GEL and DEX associated forming strings of alternating phases. Interestingly, in a small pH range close to pH 4 and pH 11, the GEL phase formed a continuous layer around the DEX droplets with a thickness of about 2 µm. The radius of the DEX droplets varied between 7 and 12 µm.

Fig. 1

 1 Fig. 1 CLSM images (128 µm x 128 µm) of emulsions formed by mixing DEX, PEO and fish GEL at different pH. The GEL, PEO and DEX phases are indicated in white, grey and black, respectively.

Fig. 2

 2 Fig. 2 Angles of the DEX (black circle), PEO (green diamond) and GEL (red square) phase as a function of the pH. Solid lines are guides to the eye.

  Fig. 3 Number of charges per gram bovine (black circles) or fish skin (red circles) GEL as a function of the pH. The solid lines are guides to the eye.

Fig. 4

 4 Fig. 4 CLSM images (128 µm x 128 µm) of DEX in PEO emulsions GEL prepared at different dilutions and pH as shown in the figure. Also shown is the evolution of the morphology with time at 20°C after the emulsions had been cooled for 1 h in a refrigerator at 8 -9°C.

Fig. 5

 5 Fig. 5 CLSM images (255 µm x 255 µm) of emulsions containing 7.2 wt% PEO, 1.7 wt % DEX and different concentrations of GEL as indicated in the figure.

Fig. 7

 7 Fig.7 Viscosity of emulsions at different dilutions, as indicated in the graph, as a function of increasing (circles) and decreasing (triangles) shear rate. Filled and open symbols represent measurements during the first and second shear ramp, respectively.

Fig. 8

 8 Fig. 8 CLSM images (128 µm x 128 µm) of emulsions prepared at D = 1.5 that were cooled in a refrigerator at 8 -9 °C and subsequently further diluted at 20 °C. The effect of aging 1 and 7 days is also shown. The red and green colors corresponded to the fluorescence signal from Rhodamine B and labeled FITC-DEX, respectively. Photographs of the samples show the slow sedimentation of the droplets for the diluted samples.

Fig. 10 .

 10 Fig. 10. CLSM images (128 µm x 128 µm) of an emulsion formed by mixing XYL, AMP and GEL one week after preparation (left) and after dilution by a factor 100 (right). For the undiluted sample, the fluorescence intensity increases in the order XYL, GEL and AMP phase. After dilution AMP and XYL mix, and the GEL phase has the strongest fluorescence.

Fig

  Fig. S3 (a) G' (circles) and G" (squares) as a function of temperature while cooling (black symbol) and 412 heating (red symbol) for aqueous beef GEL solutions at pH 7. (b) Gelation (filled symbols) and melting 413 (open symbols) temperatures of aqueous beef Gel solutions at different pH 414

Fresh

  Fig.S6CLSM images (128 µm x 128 µm) of an emulsion prepared at D = 1.5 that was cooled in a refrigerator at 8 -9 °C as a function of time after dilution at 20 °C to D = 6. The red and green colors corresponded to the fluorescence signal from Rhodamine B and labeled FITC-DEX, respectively. 2.5 h after dilution, big droplets still contained DEX, but after 5 h, the DEX was homogenously distributed.

Acknowledgments

Joyce Djitsa Djapom Martiale is thanked for her contribution to the investigation of the phase diagram and the analysis of contact angles between different solutions. This work was supported by Le Mans Metropole in the framework of the BIOREGATE consortium.

Authors contributions:

Yuwen Meng: Investigation, writing -original draft;


Maria Moutkane

Lazhar Benyahia

Taco Nicolai
email: taco.nicolai@univ-lemans.fr

Taco Orcid

Nicolai

Stable protein microcapsules by crosslinking protein particles in water in water emulsions

Keywords: Microcapsule, protein, emulsion, aqueous three phase, enzyme

Stable protein microcapsules by crosslinking protein particles in water in water emulsions

Maria Moutkane, Lazhar Benyahia, Taco Nicolai

Introduction

Microcapsules consisting of a solid, usually spherical, shell surrounding a liquid can potentially be used as carriers of molecules of interest in aqueous systems [START_REF] Wang | Microspheres and microcapsules for protein drug delivery[END_REF][START_REF] Bah | Fabrication and application of complex microcapsules[END_REF][START_REF] Corrêa-Filho | Advances in the application of microcapsules as carriers of functional compounds for food products[END_REF][START_REF] Jaganathan | Protein microcapsules: preparation and applications[END_REF]. With proper design the shell may serve as protection of the molecules so that the latter are delivered at a desired location where they are released. Hydrophobic or hydrophilic molecules can be encapsulated if the shell surrounds an oil or water phase, respectively. Microcapsules may also serve as efficient thickening agents, because they have a low density compared to homogeneous microgels and therefore fill up space with less material [START_REF] Schong | Dry heating of whey proteins leads to formation of microspheres with useful functional properties[END_REF].

The methods to prepare microcapsules containing oil has been known for some time [START_REF] Jaganathan | Protein microcapsules: preparation and applications[END_REF]. It involves the formation of an oil in water (O/W) emulsion in the presence of macromolecules that spontaneously adsorb at the O/W interface and stabilize the emulsions. Subsequently, the macromolecules at the interface are induced to bind to each other thereby forming a solid shell that encapsulates whatever was dissolved in the oil phase. A more controlled method to form monodisperse microcapsules is to utilise microfluidics with oil droplets being injected into an aqueous medium together with another aqueous solution that forms a layer around the droplet and that can be gelified [START_REF] Kim | Recent Advances in Microfluidic Production of Functional Microcapsules by Multiple-Emulsion Templating[END_REF]. In principle, microcapsules formed in this manner can also serve in all aqueous systems, but this involves replacing the oil with water, which may be difficult. In addition, encapsulation of hydrophilic molecules needs to be done subsequently by letting them diffuse through the already formed shell.

Recently, two methods have been reported to form microcapsules spontaneously in all aqueous systems. One method exploits the effect of temperature on aggregation and phase separation of certain plant proteins [START_REF] Chen | Exploiting salt induced micro phase separation to form soy protein microcapsules or microgels in aqueous solution[END_REF][START_REF] Cochereau | Mechanism of the spontaneous formation of plant protein microcapsules in aqueous solution[END_REF]. First, coacervation of the proteins into spherical micro domains is induced at room temperature by changing the pH or adding salt. Subsequently, the temperature is increased, which has the effect of rendering the proteins more soluble and crosslinking them. By fine-tuning the kinetics of each process stable microcapsules can be formed with permanently crosslinked protein shells. The second method exploits ternary phase separation of two incompatible polymers and gelatin in water [START_REF] Meng | Exploiting multiple phase separation to stabilize water in water emulsions and form stable microcapsules[END_REF][START_REF] Chen | Water-in-water-in-water emulsions formed by cooling mixtures of guar, amylopectin and gelatin[END_REF]. By fine-tuning the conditions, it is possible to induce the gelatin phase to form a continuous layer around the droplets of one polymer rich phase dispersed in the continuous phase rich in the other polymer. Subsequently, the gelatin layer can be gelled by lowering the temperature or by adding an enzyme that crosslinks the gelatin. This method has the advantage that one can encapsulate molecules of interest within the microcapsules before crosslinking the shell and the thickness of the shell can be varied. However, it requires careful tuning of the conditions of the mixture and is limited to a phases containing polymers, such as gelatin, that can form a gel.

Here we propose a third method that exploits the phenomenon that in some cases particles spontaneously adsorb at the W/W interface between two aqueous phases formed by mixing two incompatible polymers [START_REF] Chen | Exploiting salt induced micro phase separation to form soy protein microcapsules or microgels in aqueous solution[END_REF][START_REF] Dickinson | Particle-based stabilization of water-in-water emulsions containing mixed biopolymers[END_REF][START_REF] Esquena | Water-in-water (W/W) emulsions[END_REF]. The phenomenon of spontaneous adsorption of particles at the oil/water interface leading to stabilization of O/W emulsions has been known for a long time [START_REF] Pickering | Pickering : emulsions[END_REF][START_REF] Ramsden | Separation of Solids in the Surface-Layers of Solutions and'Suspensions'(Observations on Surface-Membranes, Bubbles, Emulsions, and Mechanical Coagulation).--Preliminary Account[END_REF].

It occurs when it leads to a reduction of the free energy that compensates for the loss of translational entropy. The reduction of the free energy by adsorption of spherical particles with radius R is given by [START_REF] Aveyard | Emulsions stabilised solely by colloidal particles[END_REF]:

1
where  is the contact angle of the particles with the interface, which is determined by the difference between the interfacial tension of the particles with phase A ( PA ) and with phase B ( PB ): cos()=( PA - PB )/ AB . Particles spontaneously adsorb at the interface if ( PA - PB ) <  AB . More recently this phenomenon has been observed also for W/W emulsions, but the formation of a particle layer does not always lead to stable emulsions [START_REF] Dickinson | Particle-based stabilization of water-in-water emulsions containing mixed biopolymers[END_REF][START_REF] Esquena | Water-in-water (W/W) emulsions[END_REF]. It has been shown that by crosslinking the particle layer around the oil droplets a solid shell of colloidal particles can be produced [START_REF] Bollhorst | Colloidal capsules: nano-and microcapsules with colloidal particle shells[END_REF]. Our hypothesis is that the same can be done in an all aqueous system using W/W emulsions.

We will demonstrate the feasibility of this method by adding protein microgels to an emulsion that is formed by mixing pullulan (PUL) and amylopectin (AMP), which are both food grade polysaccharides. As has already been shown in the literature, physical crosslinking of the microgels can be induced by lowering the pH towards their isoionic point [START_REF] Donato | Mixtures of whey protein microgels and soluble aggregates as building blocks to control rheology and structure of acid induced cold-set gels[END_REF][START_REF] Kharlamova | Acid-induced gelation of whey protein aggregates[END_REF]. Covalent crosslinking can be induced using the enzyme transglutaminase [START_REF] De Góes-Favoni | Microbial transglutaminase: general characteristics and performance in food processing technology[END_REF]. We will show that microcapsules can be formed with both types of crosslinks, but covalent crosslinks are needed to maintain the stability of diluted microcapsules after increasing the pH back to neutral or for periods longer than a few days. With this method the protein microgels form a monolayer around the dispersed droplets and therefore the solid shell that is formed in this manner is very thin (0.3 µm) and fragile. Thicker protein particle shells can be formed by introducing a third phase that covers the dispersed droplets and that is preferred by the microgels. To this end we used here an aqueous three phase system (ATPS) consisting of poly(ethylene oxide) (PEO), dextran (DEX) and AMP with droplets of the PEO rich phase dispersed in a continuous AMP rich phase and the DEX rich phase forming a layer around the PEO droplets. Subsequently, the protein microgels, which partitioned preferentially to the DEX phase, were physically or covalently crosslinked.

Materials and methods

Materials

POE (M w = 2.0 × 10 5 g.mol -1 ), DEX (M w = 1.6 × 10 5 g.mol -1 ), AMP (M w = 1.6 × 10 8 g mol -1 ) , glucono--lactone (GDL) and rhodamine B were purchased from Sigma-Aldrich. PUL (M w = 1.6 × 10 5 g.mol -1 ) was kindly provided by Hayashibara co. DEX and PUL were used as received. AMP was purified before use by adapting a method described by [START_REF] Bello-Perez | Macromolecular features of starches determined by aqueous high-performance size exclusion chromatography[END_REF]. Briefly, a 5 wt% AMP solution was dispersed in a mixture of dimethyl sulphoxide (DMSO) and water 95:5 v/v. Insoluble material was removed by centrifugation. The AMP in the supernatant was precipitated by adding 3 volumes of ethanol and subsequently rinsed with acetone and diethyl ether. Whey protein isolate (WPI) (BiPRO®) containing 93 wt% protein was purchased from Davisco Foods International, Inc. (Le Sueur, MN, USA). The glucono delta lactone (GDL) and the enzyme transglutaminase were procured from Acros Organics (Thermo Fisher Scientific). The powders were dissolved in ultrapure water (MilliQ).

The PEO powder contained a very small weight fraction of silica particles that were removed by centrifugation at 5 × 10 4 g for 4.5 h. Protein microgels were obtained by heating a solution of WPI at concentration of 4 wt% at pH 5.9 overnight at 80 °C. The microgels were characterized as described in detail elsewhere [START_REF] Phan-Xuan | On the crucial importance of the pH for the formation and self-stabilization of protein microgels and strands[END_REF].

Dynamic light scattering measurements showed that their hydrodynamic radius (R h ) was 150 nm and centrifugation showed that approximately 65% of the WPI was transformed into MG, with the residual forming small strand-like aggregates that do not adsorb to W/W interfaces [START_REF] Phan-Xuan | On the crucial importance of the pH for the formation and self-stabilization of protein microgels and strands[END_REF].

Emulsions were prepared by adding together aqueous stock solutions of the polymers and the microgels in the required amounts, followed by mixing under a vortex. No effect was seen of the order of mixing nor of the duration of mixing. The net charge density of the microgels was reduced by adding HCl or GDL and in some cases the enzyme transglutaminase was added after which the emulsions were again mixed under a vortex. The W/W emulsions that were studied here were formed by mixing 1.4 wt% PUL and 6.4 %wt AMP or 1.1 wt% PUL and 6.8 %wt AMP resulting in emulsion of droplets of a PUL rich phase at 25 or 20 vol%, respectively, dispersed in dispersed in a continuous AMP rich phase. The most investigated three phase system was formed by mixing 0,088 %wt POE, 0.46 %wt DEX and 7.7 %wt AMP resulting 5 vol% of a PEO rich phase and 3 vol% of a DEX rich phase dispersed in and AMP rich phase. In a few cases, ATPS with different relative volume fractions were formed by adapting the relative polymer concentrations. In the following we indicate the phases by their majority component.

Methods

Confocal Laser Scanning Microscopy (CLSM) images were obtained with a Zeiss LSM800 (Carl Zeiss Microscopy GmbH) using water immersion objectives: HC×PL APO 63× (NA = 1.2) and a HC×PL APO 25× (NA = 0.7). The samples were either inserted into acrylic wells plates or hermetically-sealed concave slides covered with glass slips. The MG were physically labeled with rhodamine B at approximately 5 ppm. The fluorescence of rhodamine B was exited at 580 nm and detected between 580 and 800 nm.

Results and Discussion

Microcapsules with protein microgel monolayer shells

W/W emulsions formed by mixing aqueous solutions of pullulan (PUL) and amylopectin (AMP) in the presence of whey protein microgels have already been studied in some detail by Macedo et al. [START_REF] Machado | Stabilization of amylopectinpullulan water in water emulsions by Interacting protein particles[END_REF]. It was shown that adsorption of the microgels at the W/W interface depends on the pH. At neutral pH, at which they are strongly charged, the microgels adsorb only at relatively high polysaccharide concentrations, i.e. when the interfacial tension between the two phases is relatively high. However, closer to the isoionic point of the microgels (pI = 5.0) [START_REF] Kharlamova | The effect of aggregation into fractals or microgels on the charge density and the isoionic point of globular proteins[END_REF], they adsorbed spontaneously also at lower concentrations. This can be explained by the reduction of the electrostatic repulsion that inhibits formation of a layer at the interface. The microgels strongly partition to the PUL phase at pH 7, but progressively partition more to the AMP phase with decreasing pH.

Crosslinking the microgels can be induced by reducing the electrostatic repulsion either by decreasing the net charge density by reducing the pH towards the pI or by adding salt which screens the interaction. For this study we reduced the net charge density either by adding aliquots of an HCl solution under strong stirring or by adding GDL. GDL slowly breaks down releasing one mol H + per mol GDL enabling one to reduce the charge density in quiescent conditions. Fig. 1 shows CLSM images of emulsions containing 20 vol% of the PUL phase dispersed in the AMP phase with 2 g/L microgels and 0.294 mmol HCl per gram protein at different times after preparation. Just after mixing very small droplets of the PUL phase covered with microgels were observed. The droplets grew initially by coalescence from about 2 µm to about 10 µm, but the coalescence stopped after a few hours and the emulsion remained stable for at least one week.

Without adding HCl the coalescence continued slowly and macroscopic phase separation was observed after one day. The addition of HCl caused a decrease of the net charge density of the microgels leading to an attractive interaction between the microgels at the interface that inhibited coalescence. However, it also caused slow aggregation of the excess microgels that can be clearly seen after standing for one week. In order to increase the attraction between the microgels, slightly more HCl was added (0.313 mmol). In this case the microgels aggregated rapidly, which hindered the formation of the layer around the PUL droplets. Instead a stable network of microgels was formed that trapped the PUL droplets, see fig. 3 Exploiting acid-induced aggregation of microgels to form a W/W emulsion gel has already been explored in some detail elsewhere [START_REF] Gonzalez-Jordan | Cold gelation of water in water emulsions stabilized by protein particles[END_REF]. These results at different HCl concentrations can be explained by the strong dependence of the aggregation rate of protein aggregates on their charge density that was reported in the literature [START_REF] Kharlamova | The effect of aggregation into fractals or microgels on the charge density and the isoionic point of globular proteins[END_REF]. The transition between the system containing well-dispersed stable droplets and microgel aggregates occurred over a remarkably narrow range of added HCl considering that 0.24 mmol/g HCl is needed to reduce the net charge of the proteins by 1 unit, taking 1.75x10 4 g/mol for the molar mass of the whey protein isolate. Similar results were obtained by adding GDL. The advantage of reducing the net charge density progressively with GDL is that the microgels can form a layer around the droplets before they strongly aggregate at lower net charge densities. By fine tuning the amount of added GDL, stable emulsions could be formed without aggregating the excess microgels, see fig. 4. However, here as well the microcapsules obtained after dilution were stable only for one or at most two days. We may conclude that the attractive interaction between the microgels induced by decreasing their net charge density is sufficient to stabilise the emulsions and the diluted microcapsules for one or two days, but it is not sufficient to stabilise the latter for longer periods. In addition, the microcapsules dispersed immediately after increasing the pH back to neutral. As was mentioned in the Introduction, covalent bonds can be formed between the microgels by enzymatic crosslinking using transglutaminase. In order to speed-up the enzymatic crosslinking of the microgels at the interface we added a relatively large amount of transglutaminase (1.5 wt%).

Nevertheless, the emulsion needed to be stable for at least a few hours in order to sufficiently crosslink the microgels without significant coalescence occurring in the meantime. Therefore, we prepared the emulsion with 0.274 mmol HCl per gram protein and the emulsion was diluted one day after preparation. Fig. 5 shows that diluted microcapsules were in this case stable for at least one week. After standing one day the microcapsules were still well dispersed. After one week the microcapsules had aggregated and sedimented, but they could be easily redispersed. The covalently crosslinked microcapsules resisted increase the pH by addition of NaOH and strong shear in a vortex.

However, the shells of many capsules were deformed even before shearing. The protein microgels were fluorescently labelled.

Microcapsules with shells of variable thickness

Microcapsules with thicker shells can be formed by adding a third component that forms a separate phase that surrounds the dispersed droplets and which captures the protein microgels. As was discussed in more detail elsewhere [START_REF] Meng | Exploiting multiple phase separation to stabilize water in water emulsions and form stable microcapsules[END_REF], one phase (A) separates the two other phases (B and C) if a b the interfacial tension between B and C is larger than the sum of that between A and B and A and C.

The three phase system we used here to demonstrate the method was formed by mixing AMP, poly(ethylene oxide) (PEO) and dextran (DEX) with AMP as the continuous phase. When this system is diluted the dispersed PEO and DEX phases first mix with each other before they mix with the AMP phase. Close to the concentration where PEO and DEX mix, the DEX phase forms a layer surrounding the PEO droplets. The protein microgels that are added to the emulsion accumulate preferentially in the DEX phase. The size of the PEO droplets and the thickness of the DEX layer can be can be increased by increasing the relative volume fractions of the PEO and DEX phase, respectively, see Fig. 6. 1.5% transglutaminase that was diluted one day after preparation and imaged after a week.

The method to form microcapsules in an all aqueous system described here should be more generally applicable. Any type of particle that spontaneously adsorbs at a W/W interface and that can subsequently be crosslinked, can serve as the shell of the microcapsule. The formation of thicker shells can be applied to any three phase system, in which one phase forms a layer around dispersed droplets of another phase. The microcapsules can then be formed either by crosslinking the polymers in the layer as was shown for gelatin in ref. [START_REF] Meng | Exploiting multiple phase separation to stabilize water in water emulsions and form stable microcapsules[END_REF], or by crosslinking other ingredients that partition to that phase. Partitioning of molecules of interest towards the dispersed droplet phase can potentially be exploited to more efficiently capture them within the microcapsules. Future research will need to be done to establish how the release of whatever is trapped in the microcapsules depends on the type of shell was produced and changing external conditions.

Conclusion

Stable protein microcapsules can be formed by crosslinking protein microgels that is spontaneously form a monlayer at the surface of droplets of a pullulan rich phase dispersed in an amylopectin rich phase. Physical crosslinks are formed after reducing the charge density of the microgels by adding an acid. By fine tuning the charge density, microcapsules can be formed that resist dilution for one or two days. However, only after covalent crosslinking by adding the enzyme a transglutaminase do the microcapsules resist dilution for at least one week, increasing the pH by adding NaOH and heating to 90°C. The shells of the microcapsules are very thin (0.3 µm) and are easily deformed. Microcapsules with thicker shells can be formed by crosslinking protein microgels that partition to a dextran rich layer that surrounds droplets of a poly(ethylene oxide) rich phase dispersed in a continuous AMP rich phase. These microcapsules remain spherical even after vortex shearing. After covalent crosslinking they resist dilution for at least one week, increasing the pH by adding NaOH and heating to 90°C.

Fig. 1 Fig. 2

 12 Fig. 1 CLSM images (200 × 200 µm) of a PUL (20 vol%) in AMP emulsion containing 2 g/L protein microgels and 0.294 mmol HCl per gram protein immediately after preparation (a), after 1 h (b), after 1

Fig. 3

 3 Fig. 3 CLSM images (200 × 200 µm) of a PUL(20 vol%) in AMP emulsion containing 2 g/L protein microgels and 0.313 mmol HCl one week after preparation. The protein microgels were fluorescently labelled.

Fig. 4

 4 Fig. 4 CLSM images (200 × 200 µm) of PUL (25 vol%) in AMP emulsions containing 2 g/L protein microgels with 0.346 mmol GDL per gram protein. The images were taken just after mixing (a), after 4h (b) and after 1 day (c). Image d was taken of the emulsion that was diluted by a factor of 20 one day after adding the GDL. The protein microgels were fluorescently labelled.

Fig. 4

 4 Fig. 4 CLSM images (200 × 200 µm) of a PUL (25 vol%) in AMP emulsion containing 2 g/L protein microgels, 0.274 mmol HCl per gram protein and 1.5% transglutaminase that was diluted by a factor of 4 one day after preparation. The images were taken one day (a) and one week after dilution (b).

Fig. 6 .

 6 Fig. 6. CLSM images (200 × 200 µm) of an ATPS formed by a PEO rich phase dispersed in a AMP rich phase surrounded by a layer of a DEX rich phase containing 4.8 g/L protein microgels at different PEO/DEX/AMP phase volume fractions: 5/3/92 (a), 11/8/81 (b), 4/29/67 (c).

Authors contributions:

Maria Moutkane: Investigation; reviewing & editing Lazhar Benyahia: reviewing & editing Taco Nicolai: Supervision, conceptualization, data curation, original draft This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.


Gireeshkumar Balakrishnan
email: gireeshkumar.balakrishnan_nair@univ-lemans.fr

Fabienne Lagarde
email: fabienne.lagarde@univ-lemans.fr

Christophe Chassenieux
email: christophe.chassenieux@univ-lemans.fr

Taco Nicolai
email: taco.nicolai@univ-lemans.fr

Quantification of very Low Concentrations of Colloids with Light Scattering Applied to Micro(Nano)Plastics in Seawater

Keywords: nanoplastics, microplastics, seawater, colloids, static light scattering, dynamic light scattering

The detection and quantification of micro(nano)plastics in the marine environment are essential requirements to understand the full impacts of plastic pollution on the ecosystem and human health. Here, static light scattering (SLS) and dynamic (DLS) light scattering techniques are assessed for their capacity to detect colloidal particles with diameters between d = 0.1 and 0.8 µm at very low concentrations in seawater. The detection limit of the apparatus was determined using model monodisperse spherical polystyrene latex particles with diameters of 0.2 µm and 0.5 µm. It is shown that the concentration and size of colloids can be determined down to about 10 -6 g/L. Light scattering measurements on seawater obtained from different locations in Western Europe show that colloidal particles were detected with DLS in seawater filtered through 0.8 µm pore size filters. The concentration of these particles was not higher than 1 µg/L, with an average diameter of about 0.6 µm. We stress that these particles are not necessarily plastic. No particles were detected after filtration through 0.45 µm pore size filters.

Introduction

The fate of plastics that end up in the sea is currently attracting much attention [START_REF] Da Costa | Nano)plastics in the environmental sources, fates and effects[END_REF]. When discharged in the environment, plastics undergo mechanical (erosion, abrasion), chemical (photo-oxidation under UV radiation, hydrolysis), and biological (degradation by microorganisms) actions [START_REF] Da Costa | Nano)plastics in the environmental sources, fates and effects[END_REF][START_REF] Andrady | The plastic in microplastics: A review[END_REF][START_REF] Barnes | Accumulation and fragmentation of plastic debris in global environments[END_REF][START_REF] Koelmans | Nanoplastics in the Aquatic Environment. Critical Review[END_REF][START_REF] Eriksen | Plastic pollution in the world's oceans: More than 5 trillion plastic pieces weighing over 250,000 tons afloat at sea[END_REF][START_REF] Moore | Synthetic polymers in the marine environment: A rapidly increasing, long-term threat[END_REF], which leads to aging and fragmentation of macroplastics into microplastics, defined as plastic particles smaller than 5 mm [START_REF]Proceedings of the International Research Workshop on the Occurrence, Effects, and Fate of Microplastic Marine Debris[END_REF]. Microplastics were found to be ubiquitous in the environment, in particular on the surface of the oceans. One important issue that has recently emerged is whether microplastics continue to fragment into colloidal particles with a diameter d < 1 µm that are often called nanoplastics [START_REF] Gigault | Marine plastic litter: The unanalyzed nano-fraction[END_REF][START_REF] Gigault | Current opinion: What is a nanoplastic?[END_REF][START_REF] Browne | Microplastic-An emerging contaminant of potential concern?[END_REF]. However, some authors consider plastic particles to be nanoplastics provided their diameter is less than 100 nm or use the expression micro(nano)plastics [START_REF] Hammer | Plastics in the Marine Environment: The Dark Side of a Modern Gift[END_REF][START_REF] Qu | Organismal response to micro(nano)plastics at environmentally relevant concentrations: Toxicity and the underlying mechanisms[END_REF]. For simplicity, we will call the particles with d < 1 µm that were detected in this study nanoplastics.

Gigault et al. [START_REF] Gigault | Marine plastic litter: The unanalyzed nano-fraction[END_REF] investigated the release of nanoplastics under UV light from weathered polyethylene and polypropylene fragments sampled from the environment. They observed that nanoplastics with a broad range of sizes were produced over a period of weeks. More recently, Ter Halle et al. [START_REF] Ter Halle | Nanoplastic in the North Atlantic subtropical gyre[END_REF] investigated seawater collected near the surface of the North Atlantic subtropical gyre. The seawater was filtered through 1.2 µm pore size filters, and the filtrate was inspected for the presence of nanoplastics with dynamic light scattering. Particles with diameters between 1 nm and 1 µm were detected in seawater that was concentrated by a factor of 200. It was suggested that these particles were mostly nanoplastics formed by the degradation of microplastics, but the authors did not provide an estimate of the concentration of nanoplastics in the seawater.

The presence of such small particles raises questions about their environmental concentration and their potential accumulation in the trophic chain. Indeed, due to their small size and specific properties, nanoplastics can be ingested by a very large range of aquatic organisms and can interact with membranes and cells [START_REF] Koelmans | Nanoplastics in the Aquatic Environment. Critical Review[END_REF][START_REF] Qu | Organismal response to micro(nano)plastics at environmentally relevant concentrations: Toxicity and the underlying mechanisms[END_REF]. Nanoplastics dispersed in the seawater could be part of the "lost plastic" that has been dumped in the sea but is no longer observed at the surface [START_REF] Eriksen | Plastic pollution in the world's oceans: More than 5 trillion plastic pieces weighing over 250,000 tons afloat at sea[END_REF][START_REF] Koelmans | All is not lost: deriving a top-down mass budget of plastic at sea[END_REF][START_REF] Thompson | Lost at sea: Where is all the plastic[END_REF]. Detection and quantification of nanoplastics in all aquatic compartments are, therefore, urgent needs. A major difficulty is that even though the total amount of plastic in the sea is huge, the concentration of nanoplastics in seawater is still expected to be very low.

Recently, various techniques have been developed to detect nanoplastics in the natural environment [START_REF] Ivleva | Chemical analysis of microplastics and nanoplastics: Challenges, advanced methods, and perspectives[END_REF]. One method involves fluorescently-labeling the nanoplastics [START_REF] Erni-Cassola | Lost, but found with Nile red: A novel method for detecting and quantifying small microplastics (1 mm to 20 µm) in environmental samples[END_REF][START_REF] Gagné | Detection of polystyrene nanoplastics in biological tissues with a fluorescent molecular rotor probe[END_REF] in order to detect them with a microscope. This method is easy to perform and can be used to measure the average size and shape of the particles. However, in most cases, the binding between the dye and plastic is not covalent, and hence leaching of the dye occurs [START_REF] Catarino | Use of fluorescent-labelled nanoplastics (NPs) to demonstrate NP absorption is inconclusive without adequate controls[END_REF]. One common dye used for labeling plastic is Nile Red [START_REF] Cole | A novel method for preparing microplastic fibers[END_REF], which generates crystals in water that lead to false identification as plastic particles. Another method that has been widely used in the literature is gas chromatography coupled with mass spectroscopy after pyrolysis [START_REF] Ter Halle | Nanoplastic in the North Atlantic subtropical gyre[END_REF]. The advantage of this method is that the limit of detection of plastics is down to a few µg/L. The main limitation of this method is the interference from natural organic matter in the aquatic environment [START_REF] Fabbri | Use of pyrolysis-gas chromatography/mass spectrometry to study environmental pollution caused by synthetic polymers: A case study: The Ravenna Lagoon[END_REF]. Another promising strategy is the use of AF4 combined with multi-angle light scattering for the detection of nanoplastics in food matrices such as fish [START_REF] Correia | Detection of nanoplastics in food by asymmetric flow field-flow fractionation coupled to multi-angle light scattering: Possibilities, challenges and analytical limitations[END_REF]. It was reported that PS nanoplastics down to a concentration of 52 µg/g of fish could be detected, but no detection was possible for PE particles. This shows that the method developed for PS cannot be applied to other types of plastics. Yet another strategy is the combination of optical tweezers with Raman spectroscopy for the detection of plastic particles with sizes between 50 nm and 20 µm [START_REF] Gillibert | Raman Tweezers for Small Microplastics and Nanoplastics Identification in Seawater[END_REF]. This method can be used to detect and isolate nanoplastics among natural organic and mineral particles in the aquatic environment, but their quantification remains difficult.

Static and dynamic light scattering techniques have the potential to yield both the average size and the concentration of colloidal particles, even if they are present at very low concentrations. The aim of the investigation reported here was twofold. First, we critically assess the potential of these light scattering techniques to quantify the concentration and size distribution of model nanoplastics in the form of polystyrene latex particles with d < 0.8 µm dispersed in water. Then we will discuss light scattering measurements on seawater sampled at different places near the coast of Western Europe. We show that the concentration of colloids with 0.2 < d < 0.45 µm is less than 1 µg/L and cannot be characterized by light scattering. The concentration of colloids with 0.45 < d < 0.8 µm is approximately 1 µg/L and can be characterized if care is taken. Of course, colloids that are detected in seawater do not necessarily consist of plastic, as mineral colloids are expected to be present. We will mention in the Discussion section how the light scattering results depend on the type of material.

Materials and Methods

Materials

Model polystyrene latex particles with diameters of d = 0.2 µm (Thermofisher, catalog number: 5020A) and d = 0.5 µm (Polysciences, catalog number: 15700) were used as received. The density of the polystyrene particles was given by the provider as 1.05 g/mL. The solid content of the latex suspensions was 10 wt% and 2.5 wt% for d = 0.2 µm and d = 0.5 µm, respectively. Particle suspensions were prepared by dilution with salt-free Milli-Q water at different concentrations: C = 10 -3 , 10 -4 , 10 -5 , and 10 -6 g/L. These standard PS latex particle surfaces are modified with a carboxylate group, which is introduced during their synthesis. The negative surface charge on the particles makes them easily dispersible in Millipore water and also makes them stable against aggregation. Neither agglomeration nor sticking of the particles to the container walls were observed.

Samples of seawater (about 2 L) were taken by hand near the surface at different locations near the coast of France, Spain, and the Netherlands: Roscoff (48°43′35.9″ N 3°58′57.0″ W), Challans (47°03′49.4″ N 2°00′41.2″ W), Toulon (43°07′15.8″ N 5°55′28.1″ E), Lanzarote (28°57′23.8″ N 13°33′16.6″ W), and Wassenaar (52°08′55.5″ N 4°19′46.6″ E) in plastic or glass bottles that had been extensively rinsed with the same seawater. At one of these locations (Toulon), seawater was taken both near the surface and, by divers, at a depth of 10 m. The samples were filtered through Acrodisc nylon membrane filters with pore sizes of 0.45 µm or MF-Millipore cellulose ester membrane filters with pore sizes of 0.8 µm. No colloidal particles could be detected by light scattering (DLS) in Milli-Q water filtered through 0.45 µm pore size filters. When colloid free Milli-Q water was filtered through the 0.8 µm filters, colloidal particles were detected with DLS, showing that these filters released particles. Therefore, it was necessary to wash the filters by filtering about 50 mL of Milli-Q water until the release of particles was no longer detected by light scattering. A number of other commercial filters were tested (Acrodisc glass membrane filters (1 µm), Whatman poly (ether sulphone) membrane filters (0.8 µm), and Whatman glass fiber filters (1.5 µm)), but they released more particles and were therefore discarded. We tested the retention of particles smaller than the pore size by comparing the scattering intensity of latex particles before and after filtration and found it to be negligible. In addition, we did not find that the scattering intensity decreased further if filtered solutions were filtered a second time.

Light Scattering

The theory of static and dynamic light scattering is briefly reviewed here. For more details, see Refs. [START_REF] Schurtenberger | Characterization of biological and environmental particles using static and dynamic light scattering[END_REF][START_REF] Filella | Analytical applications of photon correlation spectroscopy for size distribution measurements of natural colloidal suspensions: Capabilities and limitations[END_REF][START_REF] Van De Hulst | Light Scattering by Small Particles[END_REF][START_REF]The Method and Some Applications[END_REF][START_REF] Berne | Dynamic Light Scattering with Applications to Chemistry[END_REF][START_REF] Jonasz | Light Scattering by Particles in Water: Theoretical and Experimental Foundations[END_REF]. In static light scattering, the average scattering intensity of the scattering objects is measured for a given interval of time. This average scattering intensity is often expressed in terms of the Rayleigh ratio ( ) which is calculated as the average excess scattering intensity of the samples over that of the solvent normalized by the scattering of a standard, for which we used toluene with Rayleigh factor of Rref = 1.35 × 10 -5 cm -1 . Using the Rayleigh-Gans approximation, one can relate to the average molecular weight (Mw) and the structure factor of the scattering objects by the following equation (see supplementary information for details).

(

) 1
where K (K =

) is an optical constant that depends on the refractive index increment (δn/δc) and the wavelength of the light (λ). C is the concentration of the scattering objects and Na is Avogadro's number. S(q) describes the dependence of the scattered intensity on the scattering wave vector q, which itself is a function of the angle of observation θ (

). For mono-disperse spherical particles of diameter d:

(2)

For particles of any shape, the initial q-dependence of S(q) can be expressed as a series expansion in terms of the z-average radius of gyration (Rg):

(3) With DLS, one determines the correlation between the intensity at a given time with that at a delay time (t) later. The average over many starting times yields the normalized autocorrelation function of the scattered light intensity . g2(t) is related to the normalized electric field autocorrelation function (g1(t)) through the so-called Siegert relation:

(4)
The pre-factor β is smaller than unity and depends on the optical set-up. The second term in Equation ( 4) reflects the fluctuation in the number of particles (N) that are present in the scattering volume. The g1(t) obtained from dynamic light scattering measurements, could be described by a monomodal relaxation time distribution (A(τ)), which was determined by fitting the correlation functions to a general exponential function:

τ (5) with τ ( 6
)
where is a normalization constant, τgex is a characteristic relaxation time and p and s are parameters that allow different shapes of the distribution. The average relaxation rate ( = <1/τ>) is related to the z-average diffusion coefficient (D) of the particles in the suspension according to:

From the diffusion coefficient, the z-average hydrodynamic diameter (dh) of the particles was calculated using the Stokes-Einstein relation: [START_REF] Gigault | Marine plastic litter: The unanalyzed nano-fraction[END_REF] where k is the Boltzman constant and η the viscosity of the solvent.

Static and dynamic light scattering measurements were conducted using a commercial apparatus ALV/CGS3 (ALV-Langen, Germany). The light source was a He-Ne laser with wavelength λ = 632 nm. The temperature was controlled by a thermostat bath to 20 ± 0.2 °C. Measurements were made at angles of observation (θ) between 13 and 150 degrees, which correspond to scattering vectors q (ranging from 3.0 × 10 6 up to 2.5 × 10 7 m -1 . Intensity autocorrelation functions were obtained using a digital multi-tau correlator.

Results

Model Particles

We tested the limitations for static and dynamic light scattering measurements of the equipment used in this study with monodisperse polystyrene latex particles with d = 0.2 µm and d = 0.5 µm. Figure 1 shows Rθ as a function of q for the aqueous latex suspensions at C = 10 -3 , 10 -4 , 10 -5 , and 10 -6 g/L. For comparison, we also show the results for pure Milli-Q water. The dashed lines in Figure 1 represent fits to Equations ( 1) and ( 2) with d = 0.2 µm and d = 0.50 µm, whereas the solid lines represent fits to the Mie theory [START_REF] Van De Hulst | Light Scattering by Small Particles[END_REF]. The Rayleigh-Gans approximation (Equation ( 1)) gave similar results for the smaller particles, but the Mie theory described the experimental results better for the larger particles at higher q-values. The experimental value of Rθ for pure water found in this investigation is in good agreement with the value reported in the literature [START_REF] Morel | Optical properties of pure water and pure sea water[END_REF] and is shown for comparison in Figure 1a. It is important to realize that, as a consequence of the steep decrease of Rθ with increasing q for q > d -1 , suspensions of the smaller particles actually scatter more light for q > 2 × 10 7 m -1 (θ > 70°) than those of the larger particles at the same concentration, see Figure 1. The scattering intensity of the smaller latex suspension was much larger than that of water down to C = 10 -5 g/L over the whole accessible q-range. However, Rθ of the suspension of the larger latex particles at C = 10 -5 g/L q ( m -1 ) approached that of water at the highest q-values. As a general feature, the scattering intensity by suspensions of homogeneous spherical particles at a fixed mass concentration increases with increasing size for q.d << 1, but decreases for q.d > 1 as can be clearly seen from Figure 1. At a given value of q and C, Rθ is largest for particles with d  2π/q. It is, therefore, necessary to do light scattering measurements at small q-values if very low concentrations of large particles are investigated. This is illustrated here for particles with d = 0.5 µm for which the scattering intensity is close to that of water at C ≤ 10 -5 g/L if q > 2 × 10 7 m -1 , i.e., if θ > 70°, but at smaller angles they still scatter orders of magnitude more light than water even at C = 10 -6 g/L, see Figure 1b. As was mentioned above, if the number of particles in the scattering volume (N) is not large, one needs to consider the fluctuation of N in time due to the diffusion of particles in and out of the scattering volume (see Equation ( 4)). This effect can be clearly seen from the time dependence of the intensity at different concentrations. Figure 2 shows examples of Rθ as a function of time for the larger latex particles at C = 10 -4 , 10 -5 , and 10 -6 g/L at q = 3.9 × 10 6 m -1 (θ = 17°). For each solution, three measurements were conducted for a duration of 15 min. The average value of Rθ decreases in proportion to the concentration, but slow fluctuations became significant for C = 10 -5 g/L and were more important for C = 10 -6 g/L. The scattering volume of the apparatus used here was approximately 0.3 mm 3 . The average number of particles in this volume was 45 at C = 10 -5 g/L and less than 5 at C = 10 -6 g/L. It takes the latex particles about 10 3 s to diffuse 0.1 mm, which explains why the fluctuations in Rθ are very slow. As a consequence, one needs to average over very long time periods to obtain accurate averages. g/L at q = 3.9 × 10 6 m -1 (θ = 17°). Three measurements are shown for each concentration.

10 7 R   cm -1
Dynamic light scattering measurements could not be conducted reliably for suspensions of the larger latex particles at C = 10 -6 g/L because the average number of particles in the scattering volume was too low. Figure 3a shows intensity autocorrelation functions obtained at different scattering vectors for latex particles with d = 0.5 µm at C = 10 -5 g/L. The correlation functions were analyzed using Equation ( 5), assuming a log-normal size distribution. The solid lines in Figure 3a represent the fit results, and the corresponding size distributions are shown in Figure 3b. The q-dependence of the z-average hydrodynamic diameter is shown as an inset in Figure 3b. Even at this low concentration, the dh values found with DLS were within 20% of the nominal value at low q-values and within 40% at high q-values. The lower precision at higher q-values was caused by the low scattering intensity; see Figure 1. Notice that the correlation functions shown in Figure 3a did not all reach zero, which was due to the slow fluctuation of the number of particles in the scattering volume discussed above that causes an additional slow relaxation time at very low particle concentrations. This problem was greatly exacerbated at C = 10 -6 g/L and is the reason why no reliable DLS results could be obtained for that system. The solid lines represent the fit results to Equation ( 5). (b) Distributions of the hydrodynamic diameter corresponding to the fit results shown in Figure 3a. The inset shows the z-average hydrodynamic diameter as a function of q.

Colloidal Particles in Seawater

The capacity to detect and characterize colloidal nanoplastics in seawater was tested by investigating samples of seawater. The seawater was filtered through 0.8 µm or 0.45 µm pore size filters in order to assess the presence of particles smaller than 0.8 µm and smaller than 0.45 µm separately. This is necessary because the presence of a small number of large particles can hide the light scattering signal from small particles. Figure 4 shows the q-dependence of Rθ in comparison with that of Milli-Q water to which sea salt was added at the concentration found in the sea. As expected, adding sea salt caused a small increase in the scattering intensity with respect to pure water, as shown in Figure 1 [START_REF] Morel | Optical properties of pure water and pure sea water[END_REF].

Figure 4.

Dependence of the Rayleigh ratio on the scattering wave vector for seawater filtered using different pore sizes. For comparison, the results for Milli-Q water with added sea salt are also shown. The solid lines represent a fit to Equations ( 1) and ( 2) with d = 0.6 µm. Black circle for salted Milli-Q water, triangle down for filtered seawater (0.45 µm filter), and a green square and yellow diamond for filtered seawater (0.8 µm filter) collected near the surface and at a depth of 10 m, respectively.

The amount of light scattered by seawater filtered through 0.45 µm pores was within the experimental error, the same as for Milli-Q water with the right amount of sea salt over the whole q-t( µ s ) range. Considering results obtained on model latex particles discussed in the previous section, such a result would be obtained for suspensions of latex particles with d = 0.2 µm only if C < 1 µg/L and for particles with d = 0.5 µm only if C < 0.1 µg/L. The implication is that the concentration of particles in this sample of seawater with d between 0.2 and 0.45 µm was less than 1 µg/L, where we assume that the refractive index increment and the density of the particles are close to those of polystyrene, which is the case for most types of nanoplastics. Of course, this does not exclude the possibility that smaller particles are present in higher concentrations. For instance, the excess scattering intensity of very dilute suspensions of particles with d = 20 nm is a thousand times less than for d = 0.2 µm at the same mass concentration. The time-averaged value of Rθ of seawater filtered through 0.8 µm pores was much larger and decreased strongly with increasing q, which shows that it was dominated by the scattering from large particles. The solid line through the data represents a fit to monodisperse spherical particles with d = 0.6 µm. The deviation at q > 10 7 m -1 means that the particles are not monodisperse spheres, as might be expected. Comparison with the results obtained for the latex particles with d = 0.5 µm shows that the concentration of particles in the seawater with d between 0.45 and 0.8 µm was less than 1 µg/L, assuming that their refractive index and density are close to those of polystyrene. Results obtained with seawater sampled at different locations and at different depths were similar.

Figure 5 shows the scattering intensity as a function of time for seawater filtered through 0.8 µm pores taken at the surface and at a depth of 10 m. Rθ fluctuated slowly with time, indicating that the number of particles in the scattering volume was not large, as was discussed above. However, the average value of Rθ was the same at the two different depths. Results obtained with seawater samples taken at other locations were similar. Autocorrelation functions of seawater filtered with a pore size of 0.45 µm did not show significant relaxation with (g2(t) -1)  0 for t > 1 µs. This is expected as the scattering by seawater is caused by density fluctuations and the diffusion of ions, which relax on timescales shorter than 1 µs. Figure 6a shows examples of normalized intensity autocorrelation functions obtained at different scattering angles for seawater filtered with a pore size of 0.8 µm. Notice that results obtained at higher scattering angles were not trustworthy because the scattering intensity was close to that of seawater; see Figure 4. The correlation functions show a well-defined fast decay followed by an ill-defined slow decay. The fast decay is due to the diffusion of particles, whereas the slow decay is caused by fluctuations in the number of particles in the scattering volume. The fast decay was analyzed in terms of a relaxation time distribution that was converted into a distribution of dh. The fit results are shown as solid lines in Figure 6a, and the corresponding size distributions are shown in Figure 6b. In most cases, z-average hydrodynamic diameters between 0.6 and 0.8 µm were obtained, consistent with the diameter obtained from fitting the structure factor (0.6 µm). The relatively weak dependence of dh on q implies that the particles were roughly spherical and not very polydisperse, but the structure factor shows that they are not perfect monodisperse spheres either. Notice that the size distribution extends to sizes larger than the pore size. The reason is that the analysis method gives a distribution of sizes even if the particles are monodisperse. This can be clearly seen from the results on monodisperse latex particles shown in the previous section. The average diameter does, however, correspond to the true average diameter. A second reason why the distribution extends to larger values than the nominal pore size is that the 0.8 µm filters contain a distribution of pore sizes and may therefore allow some larger particles to pass.

Discussion

It was demonstrated here that it is possible to quantitatively characterize colloidal particles in aqueous suspension with static and dynamic light scattering as long as they scatter significantly more than water and the scattering volume contains at least a few tens of particles. These limitations depend on the size, shape, polydispersity, and refractive index increment of the colloids. Monodisperse spherical latex particles with d = 0.2 µm could be reliably characterized by static light scattering down to C = 10 µg/L. Latex particles with d = 0.5 µm could be characterized by static light scattering measurements down to C = 1 µg/L, but only down to C = 10 µg/L by DLS. Of course, there is no sharp boundary between concentrations that can and cannot be characterized by light scattering techniques. It is simply the case that the results become progressively less reliable when the concentration decreases.

For samples of seawater, we found that the concentration of colloids with diameters between 0.2 and 0.8 µm, was not more than 1 µg/L, assuming that they have the same refractive index increment and density as polystyrene. This concentration was barely sufficient for quantitative characterization by light scattering. The scattering of seawater filtered through 0.45 µm pores was within the experimental error, the same as that of salted water. This means that the intensity detected for seawater filtered through 0.8 µm pore size filters is due to scattering by colloids with diameters between 0.45 µm and approximately 0.8 µm.

If we consider that the amount of "lost plastic", which is estimated at about 10 14 g [START_REF] Eriksen | Plastic pollution in the world's oceans: More than 5 trillion plastic pieces weighing over 250,000 tons afloat at sea[END_REF], is distributed equally in the form of colloids in the oceans, which have a total volume of about 10 21 L, the expected concentration of nanoplastics is at most 0.1 µg/L, which was shown here to be below the limit of detection by light scattering techniques. We did not observe major differences in the amount of larger colloids in the seawater samples taken at different locations. However, these samples were all taken near the coast of Europe and may therefore not be representative of the global average concentration. On the other hand, Erikson et al. [START_REF] Eriksen | Plastic pollution in the world's oceans: More than 5 trillion plastic pieces weighing over 250,000 tons afloat at sea[END_REF] found that the distribution of microplastics (between 0.33 and 1 mm) in the North Atlantic was within a factor of 2 the same as in the other oceans. In addition, some of the samples presented here were taken in the Mediterranean Sea, which is known to be a hotspot for plastic pollution [START_REF] Cozar | Plastic accumulation in the Mediterranean Sea[END_REF]. More measurements of the concentration of colloids at different locations and depths are needed to determine their actual distribution in the oceans.

The formation of larger microplastics due to plastic fragmentation in the marine environment is well established, and its adverse effects on aquatic organisms are a serious concern. However, we still lack data to support the formation of nanoplastics with d < 1 µm in the marine environment. If we assume that these particles are formed, then the question is whether they are stable in the marine environment. It is most likely that the particles are charged when their size is < 1 µm and hence the presence of salt will induce aggregation due to screening of the surface charge of the particles [START_REF] Singh | Understanding the stability of nanoplastics in aqueous environments: Effect of ionic strength, temperature, dissolved organic matter, clay, and heavy metals[END_REF][START_REF] Tallec | Surface functionalization determines behavior of nanoplastic solutions in model aquatic environments[END_REF]. Interestingly, the presence of organic matter does not prevent the aggregation of PS nanoplastics in seawater (see references 21 and 22). In addition, it was reported that in the presence of salt, UV irradiation induces the aggregation of nanoplastics [START_REF] Li | UV/ozone induced physicochemical transformations of polystyrene nanoparticles and their aggregation tendency and kinetics with natural organic matter in aqueous systems[END_REF][START_REF] Liu | Aggregation kinetics of UV irradiated nanoplastics in aquatic environments[END_REF]. Hence, most likely, in the marine environment, nanoplastics are in aggregated form.

Unfortunately, DLS cannot inform us about the chemical composition of the detected particles. It is therefore not possible to determine whether the detected colloids are actually nanoplastics. One also needs to consider that there are many natural sources of colloids in the ocean [START_REF]Environmental Colloids and Particles: Behaviour, Separation and Characterisation[END_REF][START_REF] Delay | Nanoparticles in aquatic systems[END_REF][START_REF] Linders | Particle sources and transport in stratified Nordic coastal seas in the Anthropocene[END_REF]. Kioke et al. [START_REF] Koike | Correction: Role of sub-micrometre particles in the ocean[END_REF] reported the presence of large numbers (10 7 per ml) of submicron detrital particles in the open ocean. They found that 95% of those particles are between 0.32 and 0.6 µm and these particles were produced by the activity of small flagellates. These particles account for almost 10% of the total dissolved organic materials in the ocean.

Interestingly, it has been reported that colloidal particles form spontaneously within hours or days in seawater that was filtered through 0.45 µm or 0.22 µm pore size filters [START_REF] Chin | Spontaneous assembly of marine dissolved organic matter into polymer gels[END_REF][START_REF] Wu | Formation and characteristics of biomimetic mineralo-organic particles in natural surface water[END_REF], which was attributed either to the association of dissolved organic matter into polymer gel particles [START_REF] Chin | Spontaneous assembly of marine dissolved organic matter into polymer gels[END_REF] or to the spontaneous formation of mineral-organic particles [START_REF] Wu | Formation and characteristics of biomimetic mineralo-organic particles in natural surface water[END_REF]. We have tested whether colloids were formed in the filtered seawater samples studied here with time for up to two weeks, but we did not observe that the scattering intensity increased in any of the filtered seawater samples that were collected for this study. A possible explanation is that the glassware used in the studies reported in the literature slowly released colloidal particles. We have ourselves noted this in the past.

As mentioned in the introduction section, Ter Halle et al. [START_REF] Ter Halle | Nanoplastic in the North Atlantic subtropical gyre[END_REF] address the issue of the identification of nanoplastic particles from the large source of colloidal particles present in the ocean. They used different DLS equipment that allowed measurements only at a single high scattering angle (θ = 170°, q = 2.5.10 7 m -1 ). As we showed above, at this q-value, the scattering intensity was very close to that of seawater itself, and it was not possible to characterize the particles by DLS directly in seawater at θ = 170°. Therefore, Ter Halle et al. concentrated 1 L of seawater by a factor of 200 using ultrafiltration, which allowed them to detect colloidal particles in the seawater using light scattering at θ = 170°, similar to those shown in Figure 3. However, these authors did not perform static light scattering measurements and were therefore not able to quantitatively estimate the concentration of colloidal particles. The authors claimed that these detected particles were nanoplastics using gas chromatography combined with mass spectroscopy after pyrolysis. However, with static light scattering measurements, we show that the concentration of total colloids between diameters 0.2 and 0.8 µm is not more than 1 µg/l. This extremely low concentration makes it difficult to identify the colloids with the method used by Ter Halle et al. [START_REF] Ter Halle | Nanoplastic in the North Atlantic subtropical gyre[END_REF].

It is likely that the colloids that were detected in the seawater samples studied here were not all nanoplastics. Therefore, we need to consider how light scattering results depend on the type of material. The radius of gyration and the hydrodynamic radius do not depend on the material. However, the light scattering intensity of particles with a given size and at a given weight concentration is proportional to their density and the square of their refractive index increment. Mineral particles are denser and have a larger refractive index increment [START_REF] Babin | Light scattering properties of marine particles in coastal and open ocean waters as related to the particle mass concentration[END_REF]. Therefore, the estimated particle concentration would be even lower if it were assumed that they consisted of minerals instead of plastic.

The present study confirms that the detection and identification of nanoplastics in the environment is a very challenging research area. It would involve isolating enough colloidal particles from large quantities of seawater to allow for analysis with techniques such as Raman scattering [START_REF] Gillibert | Raman Tweezers for Small Microplastics and Nanoplastics Identification in Seawater[END_REF] and gas chromatography combined with mass spectroscopy after pyrolysis [START_REF] Ivleva | Chemical analysis of microplastics and nanoplastics: Challenges, advanced methods, and perspectives[END_REF]. The challenge is to remove all non-colloidal material and, at the same time, not introduce extraneous colloids during the isolation process.

Conclusions

The light scattering intensity of seawater samples taken at different spots off the coast of Western Europe and the Mediterranean filtered through 0.45 µm pore size filters was within the experimental error, the same as for pure water with sea salt added in the same amount as in seawater. Comparison with model colloidal particles showed that the concentration of colloidal particles with diameters between 0.2 and 0.45 µm in the seawater samples was less than 1 µg/L. Colloidal particles were detected in seawater filtered through 0.8 µm pore size filters, but the concentration was at most 1 µg/L. The dynamic light scattering measured showed that the particles had a distribution of sizes with an average hydrodynamic diameter of 0.6 µm. The concentration of colloidal particles in the seawater samples is too low to be able to characterize their composition. Measurements on model colloidal particles show that the characterization of colloidal particles in seawater requires the use of state-ofthe-art light scattering equipment that allows measurements as a function of the scattering wave vector.

Figure 1 .

 1 Figure 1. Dependence of the Rayleigh ratio on the scattering wave vector for suspensions of latex particles with d = 0.2 µm (a) or d = 0.5 µm (b) at different concentrations. The dashed and solid lines represent fits to the theory assuming the Rayleigh-Gans approximation Equation (1) and the Mie theory, respectively.

Figure 2 .

 2 Figure 2. Time dependence of Rθ for suspensions of latex particles (d = 0.5 µm) at C = 10 -4 (a), 10 -5 (b), and 10 -6 (c)g/L at q = 3.9 × 10 6 m -1 (θ = 17°). Three measurements are shown for each concentration.

Figure 3 .

 3 Figure 3. (a) Normalized intensity autocorrelation functions at different scattering wave vectors (q(m -1 )) obtained for suspensions of latex particles with d = 0.5 µm. The corresponding scattering angles are indicated in brackets. The solid lines represent the fit results to Equation (5). (b) Distributions of the hydrodynamic diameter corresponding to the fit results shown in Figure 3a. The inset shows the z-average hydrodynamic diameter as a function of q.

Figure 5 .

 5 Figure 5. Time dependence of Rθ at q = 3.9 ×10 6 m -1 (θ = 17°) for suspensions of seawater filtered through 0.8 µm pore size filters taken at the surface (a) and 10 m depth (b). Three measurements are shown for each sample.

Figure 6 .

 6 Figure 6. (a) Normalized intensity autocorrelation functions at different scattering wave vectors obtained for seawater filtered through 0.8 µm pores. The solid lines represent the fit results to Equation (5). (b) Distributions of the hydrodynamic diameter corresponding to the fit results shown in Figure 6a. The inset shows the z-average hydrodynamic diameter as a function of q.

-3 g /L 10 -4 g /L 10 -5 g /L 10 -6 g /L

Acknowledgments: François Galgani is acknowledged for providing the samples at different depths from the Mediterranean Sea.

Funding: This work was funded by Ifremer (convention 17/1212947B, project MERLIN MICROPLASTIQUES) and by the ANR CESA (ANR-15-CE34-0006-02, NANOPLASTICS project).

Institutional Review Board Statement: Not applicable.

Informed Consent Statement:

Data Availability Statement: Data available in the manuscript.

Conflicts of Interest:

The authors declare no conflicts of interest.