Marine Microgels

C H A P T E R

9 Marine Microgels Mónica V. Orellana*, Caroline Leck† *Polar Science Center, University of Washington/Institute for Systems Biology, Seattle, Washington, USA † Department of Meteorology, University of Stockholm, Stockholm, Sweden

C O N T E N T S I

Introduction 

451

II

What Are Polymer Gels? 

454

III Structure, Properties, and Dynamics of Marine Polymer Gels  A Composition of Marine Polymer Gels B Dynamics: Assembly, Size, and Stability of Gels in Seawater 1 Ionic Interactions 2 Hydrophobic Interactions  3 Size and Stability of Gels 4 Macroscopic Polymer Gels and TEP

IV Phase Transition 

454 454 456 456 458 459 463

Marine Gels in the Atmosphere and Their Relevance for Cloud Formation  A Is the “Gel Theory of Marine CCN” Coupled to the Sulfur Cycle?  B The Effect of Gels on Bubble Properties and Bursting C Is the Gel Theory for the Origin of Marine CCN Consistent with Primary Marine Aerosol Observations?

Acknowledgments 

467 467 469 471 471

References 472

464

I INTRODUCTION Marine gels are three-dimensional (3D) polymer networks proposed to play a pivotal role in regulating ocean-basin-scale biogeochemical dynamics. Microgels structurally link biological production and microbial degradative processes at the ocean’s

Biogeochemistry of Marine Dissolved Organic Matter, http://dx.doi.org/10.1016/B978-0-12-405940-5.00009-1

V

surface to biogeochemical dynamics at the ocean’s interior, cloud properties, radiative balance, and global climate (Figure 9.1). Formed in situ in the ocean’s water column, gels provide structure to the microbial loop by forming metabolic “hot spots.” Assembling from dissolved biopolymers in the microlayer at the ocean-atmosphere interface,

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Copyright © 2015 Elsevier Inc. All rights reserved.

452

9.  Marine Microgels

FIGURE 9.1  Marine polymer gel dynamics. DOM is mainly produced by phytoplankton and bacteria, which release free biopolymers and/or micron-size gels into the seawater by diverse mechanisms including disaggregation of phytoplankton cells (Table 9.1). Gel assembly is an important mechanism by which refractory short chain biopolymers (~1 nm or 1000 Da; Aluwihare et al., 1997; Chin et al., 1998) form “hot spots” for microbial activity (Azam, 1998; Azam and Malfatti, 2007; Baltar et al., 2010), which are particularly important in the dark ocean (Herndl and Reinthaler, 2013). Although microgels are non-sinking soft matter particles, they could also be exported to the deep ocean by undergoing volume phase transition which might increase their density and settling velocity (Chin et al., 1998; Orellana and Hansell, 2012). Furthermore, microgels can form cloud condensation nuclei (CCN), which have an important role in climate feedback (see text; Orellana et al., 2011b).

gels are also found in clouds, serving as cloud condensation nuclei (CCN). Problematically, the quantitative role of marine gels in these important processes is yet to be decided. The dissolved organic carbon (DOC) pool is the largest bioreactive reservoir of organic carbon (C) in the world’s oceans (662 ± 32 Gt; Hansell et al., 2009). This pool of C is similar in size to the atmospheric reservoir 750 Gt C (Sarmiento and Gruber, 2006). The sizes of these pools relative to the fluxes between them imply that changes in C flux through DOC can significantly influence the global C cycle on relatively short timescales (Hedges and Oades, 1997; Kwon et al., 2009). Scientists have made tremendous progress in understanding DOC dynamics (Hansell, 2013; Hansell and Carlson, 2002; Hansell et al., 2009); however, we still have only a fragmentary knowledge of the source and sink mechanisms, and specifically of the role played

by biopolymers and their dynamics. Marine polymer dynamics can be viewed in the context of soft matter physics, with clear benefits for developing accurate models of the response of biogeochemical cycles to environmental forcing. Chin et al. (1998) applied the principles of soft matter physics to understand marine biopolymer dynamics, demonstrating that they assemble into 3D gel networks. By applying the tools and the conceptual framework of polymer physics, a mechanistic understanding of the structure of the DOC field and of the dynamics of DOC biopolymers emerged. This perspective brought new insights and a complementary “soft-­material”related understanding to DOC; assembly has thus emerged as a dominant concept to mechanistically explain the abiotic formation of particles in seawater, known since the time of Riley (1963), Alldredge and Jackson (1995), Chin et al. (1998), Sheldon et al. (1967), and Wells (1998).



I Introduction

Assembly of marine polymer gels occurs in the oceans when a chemically heterogeneous polydispersed mixture of biopolymers interact to form randomly tangled 3D cross-linked networks held together by ionic bonds, hydrophobic forces, and/or hydrogen bonds, depending on the nature of the polymers and the relation with the solvent (in this case seawater) (Chin et al., 1998; Ding et al., 2008; Orellana et al., 2011b; Radić et al., 2011). Marine polymer assembly is reversible, with an approximate thermodynamic yield at equilibrium of at least 10% in surface waters (maybe lower in deep waters); thus, 10% of the DOC remains in dynamic and reversible assembly equilibrium, forming porous networks (Chin et al., 1998; Orellana et al., 2011b). As a result, marine polymer gels may account for at least ~70GC of the global organic DOC pool (Verdugo, 2012). This colossal mass of polymer gel represents the largest pool of biodegradable organic C available to the microbial loop (Orellana and Verdugo, 2003; Verdugo and Santschi, 2010), which might be especially important in the dark ocean (Herndl and Reinthaler, 2013). Micronsized gels can further aggregate to form macroscopic polymer gels (Radić et al., 2011; Svetličić et al., 2005). This dynamic conceptual framework constitutes a particle size continuum for dissolved-to-gel-to-particulate organic matter (POM) in seawater (Azam, 1998; Chin et al., 1998; Koike et al., 1990; Verdugo et al., 2004) that ranges from single dissolved monomers and oligomers, to nanometer-colloidal and micron-sized gel networks (nm to μm), to larger size (mm) macrogels and aggregates (Svetličić et al., 2011; Verdugo et al., 2004). The particle size continuum is a critical mechanism through which truly dissolved organic matter biopolymers (<10 nm; Aluwihare et al., 2005; Chin et al., 1998; Jiao et al., 2010), characterized by being old, short-chain, and biologically refractory (Walker et al., 2014), are transferred to nutrient-rich gel networks, creating discrete “hot spots” accessible to microbial degradation and remineralization (Figure 9.1; Amon and Benner, 1994, 1996; Azam, 1998; Azam

453

and Malfatti, 2007; Orellana and Verdugo, 2003; Verdugo and Santschi, 2010; Verdugo et al., 2004). Excellent reviews of marine polymer gels (Verdugo, 2012; Verdugo et al., 2004; Verdugo and Santschi, 2010) and books already explain soft matter physics (Edwards, 1986; Grosberg and Khokhlov, 1994; Poon and Andelman, 2006). This chapter has a different aim: to comprehensively summarize the salient aspects of recent discoveries about the role polymer gels play in DOC dynamics. Some fundamental aspects of DOC dynamics are well documented (Hansell, 2013; Hansell and Carlson, 2013; Hansell et al., 2009), but polymer gel theory and relevant mathematical and modeling tools (de Gennes and Leger, 1982; Edwards, 1986; Ohmine and Tanaka, 1982; Tanaka et al., 1980) have not been systematically applied to achieve a mechanistic understanding of the dynamics of marine biopolymers, their emergent physical properties that arise from their interactions with seawater, or their control by environmental stimuli. Nor do we fully understand their role in colloidal trace metal scavenging (Wells, 2002), the microbial loop, C cycling, and the microbial pump, trace metal complexation and size reactivity relationships, or biogeopolymer condensate formation or their role in cloud formation (Orellana et al., 2011b).

FIGURE 9.2  Microgel sorted from seawater by flow cytometry. From Orellana et al., 2007 with modifications.

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9.  Marine Microgels

II  WHAT ARE POLYMER GELS? Chin et al. (1998) offer this summary: Polymer gels are a distinctive form of supramolecular organization formed by a deformable “3D polymer network and a solvent which, in the case of marine hydrogels, is seawater (Figure 9.2). Although the solvent prevents the collapse of the network, the network entraps and holds the solvent, creating a microenvironment that is in thermodynamic equilibrium with the surrounding media. The polymer chains that form the 3D network are interconnected by chemical (covalent bonds) or physical cross-links. In physical gels, polymers are interconnected by tangles and/or low energy links” (hydrogen bonding, metal coordination, hydrophobic forces, van der Waals forces, pi-pi interactions and electrostatic effects) “are continuously being made and broken, thus these networks are in continuous assembly/dispersion equilibrium.” “Depending on the characteristics of the polymer chains (such as polyelectrolytic properties, degree of hydrophobicity, length, and linear or branched chains) and the dielectric properties of the solvent, osmolarity, ionic composition, pH, and environmental conditions (temperature, pressure) the polymers in the gel’s matrix can interact strongly with each other, with the solvent, or with smaller solutes. These interactions determine the gel’s physical properties.” This range of structural and ultra-structural features give polymer gels unique emergent physical properties (assembly, volume phase transition; de Gennes, 1992; de Gennes and Leger, 1982), as well as unique chemical and biological reactivities. These properties are very different from those of the dispersed polymeric components comprising the gels. The soft matter “polymer gel concept” offers a solid and powerful physicochemical predictive theory that mechanistically explains the dynamics of biopolymers in the DOC pool, thereby aiding our understanding of DOC biogeochemistry (Verdugo, 2012).

III  STRUCTURE, PROPERTIES, AND DYNAMICS OF MARINE POLYMER GELS A  Composition of Marine Polymer Gels Marine gels are tangled 3D networks that assemble from the complex and heterogeneous polydispersed mixture of biopolymers present in marine and freshwater dissolved organic matter (DOM) (Chin et al., 1998; Pace et al., 2012; Verdugo, 2012). These polymer gels range in size from colloidal (1-1000 nm) to several microns and even to meters, such as those reported for the Adriatic Sea (Svetličić et al., 2005; Verdugo et al., 2004). Fluorescent probes, histochemical stains, and fluorescently labeled antibody probes demonstrate that gels contain polyanionic polysaccharides, proteins (Table 9.2), nucleic acids, and other amphiphilic and hydrophobic moieties (Chin et al., 1998; Ding et al., 2008; Orellana et al., 2007, 2011b). This composition is not surprising given the diverse spectrum of biopolymers in DOM, though DOM itself has not been completely characterized (Aluwihare et al., 1997; Benner, 2002; Close et al., 2013; Hansman et al., 2009; Hertkorn et al., 2006; Kaiser and Benner, 2008; Kaiser and Benner, 2012; McCarthy et al., 1993, 1998; see Chapter 2). Phytoplankton and bacteria in the euphotic zone are known to produce a complex variety of polysaccharides and proteins, and their monomers (amino acids and sugars) have been isolated from the DOM pool (Aluwihare et al., 2005; Benner, 2002; Jiao et al., 2010; Kaiser and Benner, 2008; McCarthy et al., 1993). The mechanisms of production of these biopolymers include several processes (Table 9.1). Phytoplankton alone can release ~10-30% of their primary production, depending on the species (Wetz and Wheeler, 2007) and their physiological stage (Biddanda and Benner, 1997; Biersmith and Benner, 1998), into the DOC pool in the form of tangled polymers forming micron-sized gels and/or free biopolymers (Biller et al., 2014; Chin et al., 2004; Orellana et al., 2011a). While some species secrete b ­ iopolymers as



III  Structure, Properties, and Dynamics of Marine Polymer Gels

TABLE 9.1  Biopolymer Production Mechanisms by Phytoplankton, Bacteria, and Other Sources (the Relative Importance of Each is Unknown) Process

Reference

Direct release

Decho (1990)

Viral lysis

Suttle (2007); Vardi et al. (2012)

Apoptosis and programmed Berman-Frank et al. (2004); cell death Bidle and Falkowski (2004); Orellana et al. (2013) Microbial degradation of particulate matter

Nagata and Kirchman (1997)

Grazing

Strom (2008); Strom et al. (1997)

Zooplankton sloppy feeding

Jumars et al., 1989

Particle dissolution

Azam and Long (2001); Carlson (2002); Kiørboe and Jackson (2001); Smith et al. (1992); Nagata et al. (2010)

Vesicle production and regulated exocytosis

455

TABLE 9.2  Proteins Found in Marine Polymer Gels (Orellana and Hansell, 2012; Orellana et al., 2007; Powell et al., 2005) ATP/GTP-binding site motif A (P-loop) Cation channel, non-ligand-gated cation channel Eukaryotic thiol (cysteine) protease cysteine-type endopeptidase Peptidase activity, tail specific Eukaryotic thiol (cysteine) protease cysteine-type endopeptidase activity Enolase Actin/actin-like protein

Biller et al. (2014); Chin et al. (1998, 2004); Orellana et al. (2011a)

polymer gels (e.g., Phaeocystis (Chin et al., 2004), Fragilariopsis cylindrus (Aslam et al., 2012)), the ratio of polymers released as gels to free polymers is unknown for most phytoplankton species. These 3D gel networks contain polysaccharides, in line with the molecular-level analysis of DOM (Aluwihare and Repeta, 1999; Aluwihare et al., 1997; Benner, 2002; Biddanda and Benner, 1997; Kaiser and Benner, 2009; see Chapter 2); peptides, protein, including ribulose-1,5-­bisphosphate oxygenase (RuBisCO; Table 9.2; Chin et al., 1998; Orellana and Hansell, 2012; Orellana et al., 2007); hydrophobic and amphiphilic moieties including phospholipids (Orellana et al., 2013); nucleic acids (Chin et al., 1998); and other metabolites (Kujawinski et al., 2009). Bacteria also release biopolymers, and ~50% of the bacterial production in the ocean is released into the DOM pool by viral burst and mortality (Suttle, 2007), probably releasing ­ bacterial

Aldose 1-epimerase/galactose metabolism Zn-finger (putative), N-recognin ubiquitin-protein ligase activity) (serine/threonine-protein kinase) Transporting ATPase Glucan 1,4-alpha-glucosidase ATP/GTP-binding site motif A (P-loop) RuBisCO (ribulose-1,5-bisphosphate carboxylase oxygenase)

membrane porins containing d-amino acid enantiomers (d-alanine, d-glutamic, d-serine, d-­ aspartate, and d-glutamate; Benner and Kaiser, 2003; Benner et al., 1992; Kaiser and Benner, 2008; McCarthy et al., 1993; Ogawa et al., 2001; Tanoue et al., 1996; Tanoue et al., 1995), or by direct production of d-amino acids when bacteria detach from particles, similarly to the breakdown of biofilms (Kolodkin-Gal et al., 2010); hydrolyases and other exoenzymes (Arnosti, 2011; Smith et al., 1992); carbohydrates and amino sugars, such as glucosamine, galactosamine, and muramic acid (an amino sugar from peptidoglycan), a bacterial cell wall polymer (Benner and Kaiser, 2003; Kaiser and Benner, 2009; McCarthy et al., 1993); fatty acids, phospholipids, and lipopolysaccharides (Popendorf et al., 2011; Wakeham et al., 2003) perhaps produced by virus while infecting and lysing phytoplankton cells (Vardi et al., 2009). Bacteria can also release biologically refractory

456

9.  Marine Microgels

and c­ hemically unknown molecules (Jiao et al., 2010; Nagata et al., 2010). This DOM pool of microbial origin, estimated at 165 PgC (Benner and Herndl, 2011), resists degradation and is typically described as high molecular weight (HMW) DOC (>1000 Da but <0.1-0.2 μm; 1 nm pore size; Benner et al., 1992). This pool is chemically defined as containing a high proportion of acetylated polysaccharides (APS) (Aluwihare et al., 1997), composed mainly of neutral monosaccharides and amino sugars that accumulate in the deep ocean (Aluwihare et al., 2005). It probably has a distinct δ15 N-DON signature, indicating complex degradation (Calleja et al., 2013), and proves itself to form the bulk of the DOC pool; 10-30% of all DOM polymers assemble into forming gels.

B  Dynamics: Assembly, Size, and Stability of Gels in Seawater Marine gels are held together by ionic, hydrophobic, or hydrogen interactions, with the most dominant being ionic interactions. 1  Ionic Interactions Irrespective of the biopolymer chemistry, biopolymer interactions in the ocean’s DOM pool spontaneously form 3D microscopic polymer hydrogel networks that continuously assemble and disperse at equilibrium (Figure 9.3; Chin et al., 1998). These micron-sized gels form hydrated Ca2+-linked supramolecular networks whose interactions with the solvent (seawater) provide them with a gel-like texture (de Gennes and Leger, 1982). Using independent and parallel methods, including dynamic laser scattering (DLS) or photon correlation spectroscopy, flow cytometry, and environmental scanning electron microscopy, Chin et al. (1998) demonstrated that polyanionic polymers (proteins, nucleic acids, polysaccharides) associate to form microscopic polymer gels as a result of the local interactions of negatively charged polymers and multivalent cations (such as the metal ions present in sea water). When these authors chelated the metals in seawater with e­ thylenediaminetetraacetic

Dissolved to gel to particulate organic matter size continuum

1 nm

1 µm

Ca2+ 2+

Ca Assembly

Ca 2+

Ca

Dispersion

Ca2+

2+

Ca

Ca

Ca2+ Annealing

Ca2+ Fragmentation 2+

Ca2+ Ca

2+

Ca2+

Ca2+

Nanogel

Ca2+

Ca2+

Ca2+ Ca2+ Ca2+

Ca2+

Macromolecules

Ca2+ 2+

Ca2+

Ca2+ Ca2+

Microgel

FIGURE 9.3 Dynamics of self-assembling polyanionic marine polymer gels. Hydrogels consist of a three-­ dimensional polymer network. Polyanionic polymers found in the DOM pool (662 PgC) assemble spontaneously, forming nanometer-sized tangled networks that are stabilized by Ca2+ bonds. The tangled nature of these nanogels allows polymers to interpenetrate neighboring gels, annealing into larger ­microgels. Polymer gels exhibit emergent properties that are different from those of the dispersed polymers that make up these networks. Polymer chains inside the microgels interact to creating a microenvironment of high-substrate concentration serving as a rich source of substrate to microorganisms. From Orellana and Verdugo (2003), with modifications.

acid (EDTA, 10 mM) or removed them by dialysis using Ca2+-free artificial seawater, the microscopic polymer gel networks did not form, indicating the cross-­linking role of Ca2+. Using x-ray energy dispersive spectroscopy, they further demonstrated that the polymer association was driven mainly by Ca2+ ionic bonds (binding sites on marine polymers include carboxyl, hydroxyl, phosphate, sulfate, amino, and sulfhydryl groups (Benner et al., 1992; Hung et al., 2003; Hung et al., 2001; Mopper et al., 1995; Zhou et al., 1998a). This finding is consistent with Ca2+ ions attaining high concentrations in seawater (10 mM). Furthermore, Ca2+ has a smaller Stokes radius than other abundant marine divalent metal ions (e.g., Mg2+), thus increasing the probability of interaction and linking the polyanionic marine polymers. Calcium also acts as an important cross-linker



III  Structure, Properties, and Dynamics of Marine Polymer Gels

in other n ­ atural gels such as pectin, alginates, and mucous (Pollack, 2001; Verdugo, 1994). While the spontaneous assembly of polymers is common in seawater, its variability is determined by the mass and composition of the biopolymer backbone, the concentration and charge density of biopolymers (i.e., coastal vs. oceanic and temporal variability), and by the concentration, valence, size, and shape of the cross-linking counter anion (i.e., coastal vs. oceanic waters, surface vs. deep waters). For example, while Ca2+ cross-linking might dominate, trace metals, which reach extremely dilute concentrations in seawater (e.g., low nM), exhibit multivalent properties (i.e., Fe3+, Al3+), allowing the fast spontaneous assembly of very stable networks that have important consequences for biogeochemistry. In fact, Wells and Goldberg (1991, 1992) demonstrated that colloids (120 nm) in coastal California surface waters contain polycations such as Fe and Al (see below). Chin et al. (1998) also demonstrated that spontaneous assembly is reversible, follows ­second-order kinetics, and, at equilibrium, has an average thermodynamic yield of 10% at room temperature. New measurements indicate that the equilibrium yield at in situ temperature can reach 30%, as in the case of Arctic biopolymers (see below, Orellana et al., 2011b). Reversibility refers to the fact that, at equilibrium, the electrostatic interactions between the polymers continuously break and form, assembling and dispersing, so that polymers that form a gel are in dynamic equilibrium with chains that remain in the bulk solution. Second-order kinetics characterizes the system’s behavior over time as a two-step assembly. Free polymer chains with molecular dimensions (from angstroms to a few nanometers) first form nanometer-sized gel networks (100-200 nm), which then anneal and interpenetrate themselves by the mechanism of diffusional reptation (thermal motion of long linear, entangled macromolecules in concentrated polymer solutions) to form colloidal-sized gels (visualized by transmission atomic force microscopy), which subsequently anneal and

457

aggregate to form micron-sized gels that stabilize in vitro and at room temperature to about 4.5-5 μm (microgels) as measured by DLS and visualized by environmental electron microscope (Figure 9.3; Chin et al., 1998). In the oceans, with an enormous availability of free polymers, gels may assemble to yet greater sizes. Physical aggregation and agglomeration have often been used to explain marine colloidal growth into larger particles, as well as the transfer of truly dissolved substances into micron-size particles (larger than 1.0 μm in size; Paerl, 1973; Riley, 1963; Wells, 2002; Wells and Goldberg, 1993). However, the mechanism and nature of the physical interaction of the polymers was not completely understood then (Paerl, 1973; Wells and Goldberg, 1993). Referring to the discovery of self-assembly, Wells (1998) commented that the “assembly processes mechanistically explains the colloidal annealing and aggregation behavior, including trace metal colloidal complexation in seawater” (Wells, 2002). The influence of attractive interactions on the aggregation of biopolymers and gel formation in colloidal systems is now well described (Poon and Andelman, 2006). In fact, the spontaneous self-assembly of marine biopolymers is consistent with the time frames of colloidal aggregation of radioactive thorium species (e.g., 234Th, 230 Th, 228Th), a process known as “colloidal pumping” (Guo and Santschi, 1997; Guo et al., 2000; Honeyman and Santschi, 1988; Santschi et al., 1995). Th species have also been used as a tracer in a range of processes, including particle cycling (Clegg and Whitfield, 1990, 1991; Dunne et al., 1997), C export flux (e.g., Buesseler, 1998; Murray et al., 1989), boundary scavenging (Santschi et al., 1999), and paleo-circulation (Moran et al., 2002). Scavenging of dissolved iron (Fe3+), a key limiting nutrient in the high-nitrate, low-chlorophyll (HNLC) regions of the Southern Ocean, as well as in the subarctic and equatorial Pacific (Martin et al., 1990, 1994; Moore and Doney, 2007), is consistent with the conceptual model developed for the scavenging of Th isotopes, whereby particle scavenging is a two-step process of scavenging

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9.  Marine Microgels

by organic colloidal and small particulates followed by aggregation and removal on larger sinking particles. Removal of dissolved iron from subsurface waters (where iron concentrations are often well below 0.6 nM) occurs by “aggregation” and by sinking particles of Fe3+ bound to organic colloids (Moore et al., 2002). In this respect, according to polymer theory, the assembly of biopolymers is accelerated not only by the presence of metal-binding ligand groups and siderophores (Butler and Theisen, 2010) but also by the fact that electrostatic interactions in polyanionic polymers form cross-links that are proportional to the square of the valence of the counter ion (Fe3+), conferring remarkable stability to the 3D gel architecture (Ohmine and Tanaka, 1982; Okajima et al., 2012; Verdugo, 2012). Chuang et al. (2013) demonstrated the role of proteins, polysaccharides, sugars such as uronic acids (containing carbonyl and carboxylic acid functional groups), and cathecolamines as major carriers of radionuclides and other metals in the Atlantic Ocean. Cathecolamines, siderophores, and uronic acids are known to assemble as gels as well (Menyo et al., 2013; Okajima et al., 2012). Additionally, sacran, a cyanobacterial sugar containing uronic acids, easily forms gels with trivalent metal ions (Fe3+, Ga3+, Al3+) (Okajima et al., 2012). Measuring the kinetics of marine biopolymer assembly and iron complexation in conjunction with electron probe analysis would improve our mechanistic understanding of iron scavenging, as well as other ions in the oceans. 2  Hydrophobic Interactions Polymer gels also form at hydrophobic interfaces, such as at the air-water interface and at bubble surfaces (Orellana et al., 2011b; Verdugo, 2012; Wheeler, 1975). Although the bulk of the polymers produced by phytoplankton and bacteria are polyanionic, they also produce amphiphilic polymers (Decho, 1990; Orellana et al., 2007, 2011b; Stoderegger and Herndl, 2004; Wingender et al., 1999). However, the assembly of marine amphiphilic polymers is less well

understood. Ding et al. (2008) demonstrated that nanomolar concentrations of amphiphilic exopolymers (20  μg L−1) released by the proteobacteria Sagittula stellata induced polymer network formation with very rapid rates of assembly. Likewise, the remarkable predominance of amphiphilic siderophores secreted by oceanic bacteria enables microbial iron acquisition, thus playing an important role in upper-ocean iron cycling (Butler and Theisen, 2010; Martinez et al., 2003). Polymers secreted by phytoplankton, such as the cyanobacteria Synechococcus, the prymnesiophyte Emiliania huxleyi, and the centric diatom Skeletonema costatum, contain proteins exhibiting hydrophobic domains (with 30% being hydrophobic amino acids) that self-­assemble in Ca2+-free seawater or in low Ca2+ concentrations (Ding et al., 2009). While the magnitude of the production of hydrophobic moieties in the world oceans is not well known, the presence of lipids in the North Pacific subtropical gyre indicates that lipid compositional signatures of colloidal-sized particles (0.2-0.5 μm) are depth specific (surface vs. mesopelagic) and that these particles account for an important percentage of the exported material in oligotrophic waters, making an important contribution to the biological pump (Close et al., 2013). Therefore, these moieties probably play an important role in self-assembly of colloidal-sized gels, as has also been demonstrated in the Arctic. In fact, in the high Arctic, amphiphilic and hydrophobic moieties play a significant role in accelerating the assembly of gels at the water-air interface, where self-assembly takes place in a matter of minutes (Figure 9.4). Hydrophobic moieties form the core of Arctic gels, containing lipids and proteins that include hydrophobic amino acid residues such as leucine, isoleucine, phenylalanine, and cysteine (Figure 9.5; Orellana et al., 2011b). These amino acid residues create hydrophobic pockets capable of allowing the assembly and further aggregation of polymer gels (Maitra et al., 2001). Aromatic residues (tryptophan and phenylalanine) may also be important. Perhaps ­hydrophobic polymers are



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III  Structure, Properties, and Dynamics of Marine Polymer Gels 50

SML Control

Gels (µM C)

40 30 20 10 0

0

10

20

30 Time (h)

40

50

60

FIGURE 9.4  Polymer gel assembly as a function of time in high Arctic surface waters (87-88° N, 2-10° W). The assembly of polymer gels was monitored by measuring percent polymers assembled as microgels (Ding et al., 2007) at 4 °C (triangles). Control experiments in which Ca2+ was chelated from seawater with 10 mM EDTA showed no assembled gels, regardless of the time of observation (squares). Each point corresponds to the average of three replicates. Note that the assembly kinetics is very fast, with the concentration and size of assembled polymer gels reaching equilibrium in 6 h. An average yield of assembly equal to 32% of the polymers present in the DOM pool was measured for either subsurface (SSW) or surface microlayer (SML) water samples. From Orellana et al., 2011b.

more abundant in recently produced gels containing a higher concentration of p ­ roteins with amphiphilic side chains, and therefore having lower C:N ratios, during and at the end of phytoplankton blooms, as in the case of the Arctic gels. Polymers with higher nitrogen content biodegrade faster in the water column (Cherrier and Bauer, 2004; Cherrier et al., 1996; Davis and Benner, 2007) and are more abundant in the surface than deep oceans (Hopkinson and Vallino, 2005; Walker et al., 2014). Walker et al. (2014) demonstrated relations between age, C:N ratio, and size of the particles; older particles are smaller and nitrogen-poor. These relations may provide an explanation for the preferential storage of polyanionic molecules in the ocean and for their forming the bulk of the interacting polymers that spontaneously assemble into polymer gels. 3  Size and Stability of Gels Polymer length, concentration, and charge density are key features conferring stability to marine 3D gel networks. The probability of assembly, the equilibrium size, and the stability of tangled

FIGURE 9.5  Hydrophobic moi-

(a) D-lle

7 Enrichment (SML/SSW)

eties in polymer gels. (a) Several cloud microgels assembled by amphiphilic polymers and stained in red (Nile Red), indicating the presence of hydrophobic moieties, and hydrophilic moieties stained green with quinacrine. (b) Enrichment of dissolved hydrophobic amino acids (leucine, isoleucine, phenylalanine, and cysteine) in the surface microlayer (SML) with respect to subsurface (SSW) waters (Orellana et al., 2011b). x-axis is sampling day-of-year (DoY); y-axis is the observed enrichment of the amino acids (see symbol legend) in the surface microlayer relative to sea surface water.

L-lle

6

D-Leu

L-Leu

5

D-Phe

4

L-Phe

3 2 1 0

(b)

226

228

230

232

236

234 DoY

238

240

242

244

9.  Marine Microgels

gel networks are determined by polymer length (Edwards, 1986). The assembly of polymers into polymer gels and their stability once assembled increases with the square of the polymer length (de Gennes and Leger, 1982; Edwards, 1986). Longer chains assemble into more stable and bigger equilibrium size polymer networks. When the length of the polymers increases, the interactions between the polymer chains (ionic, van der Waals, hydrophobic, etc.), tangles, and interpenetrations become stronger, requiring more energy to disrupt them. In contrast, short-length chains experience lower degrees of interactions, and therefore increased gel destabilization. Using ultraviolet (UV)-B irradiation to photocleave free marine polymers and assembled microgels, Orellana and Verdugo (2003) demonstrated that as the biopolymers fragmented with increasing exposure time to UV-B (λ = 280-320 nm), the assembly kinetics of those short, fragmented polymer chains took more time to reach equilibrium and the polymer gel size at equilibrium decreased exponentially (Figure 9.6). Thus, depending on the extent of fragmentation, UV-B-irradiated polymers assembled into smaller, less stable submicron-sized gel networks; at the end (>12 h of UV exposure), monomers and short oligomers completely failed to anneal and to form stable networks. Assembled gel networks irradiated with UV-B dispersed, and the resulting shortchain polymers failed to assemble into stable gels, drastically disrupting the exchange between the dissolved and the gel phases (Figure 9.6; Orellana and Verdugo, 2003). A gel’s network dependency on polymer size can explain the distribution of biopolymers in the oceans. For example, at the end of spring blooms, phytoplankton release a high percentage of their primary production as long polysaccharide chains and proteins that generally assemble to form large gel networks, as occurs with gels produced by Phaeocystis blooms in polar regions (Janse et al., 1996; Vernet et al., 1998) and ­macroscopic-sized, gel-like transparent exopolymer particles (TEP) (Alldredge et al., 1993; Mopper et al., 1995;

6

Hydrodynamic diameter (µm)

460

4

2

0

0

2

4 6 Time of assembly (days)

8

10

FIGURE 9.6  Effect of UV-A and UV-B radiation on spontaneous assembly of DOM polymers. Assembly of polymers follows characteristic second-order kinetics, forming polymer gels that grow from colloidal (nanometer) to multi-­ micron size in ~60 h. Filled squares represent the assembly of non-irradiated polymers (the control run). Assembly of polymers irradiated for 24  h with UV-A (10  W  m−2, λ = 320–400 nm) are in open squares, showing no statistical difference between the assembly kinetics of controls and UV-A-irradiated samples. The assembly of DOC polymers in seawater samples exposed to UV-B (0.5 W m−2) for 30 min (filled diamonds), 1 h (open triangles), 6 h (filled circles), and 12 h (open circles) follow a similar second-order kinetic profile; however, the time to reach equilibrium is longer and the equilibrium size of the polymer gel networks is smaller and less stable. Data points correspond to the mean ±SD of 30 dynamic laser scattering measurements. Adapted from Orellana and Verdugo (2003).

Passow, 2002b), as well as protein-containing particles (Long and Azam, 1996). An extreme case of macroscopic accumulation (meters) of polymer networks held together by hydrogen bonds took place in the Adriatic Sea, with the polymers produced by the diatom Cylindrotheca closterium (Svetličić et al., 2005, 2011). Similarly, the high concentrations of longer younger polymers produced at the end of spring blooms perhaps explains the high yield of gels found in the high Arctic at the end of a bloom of the ubiquitous diatom Melosira arctica (30% of polymers present as DOM, assembled as gels; Orellana et al., 2011b). Polymer gel theory can also explain why the distribution of marine DOM is highly skewed



III  Structure, Properties, and Dynamics of Marine Polymer Gels

toward low molecular weight (LMW) components (Benner, 2002; see Chapter 2). Truly dissolved substances make up 70 ± 5% of the total organic C, and the HMW fraction and colloids essentially make up the remaining 25% (Benner, 2002). Because the probability of short-chain oligomers and monomers to interact and form stable gel networks is very low or nil, they accumulate and probably account for the short chain (~1  nm), biologically refractory DOC pool found in the world oceans (Druffel et al., 1998; Druffel and Williams, 1990; Walker et al., 2008, 2014). Macroscopic gels form sporadically, mainly at the end of spring blooms (Passow, 2002b) when newly long polymeric chains are released by phytoplankton (diatoms), aggregating to form big particles embedded with cells (Alldredge and Jackson, 1995), degrading and/ or sedimenting as POM relatively quickly in the water column (Figure 9.1; Smetacek, 2000). However, HMW colloids are the most abundant gels within this pool. Most importantly, some 10-30% of this pool of polymers anneal to form micron-sized 3D gel networks, representing the biggest and most important shunt of polymer material that transforms the dissolved DOC into a gel phase, and then to POM in seawater (Verdugo, 2012; Verdugo et al., 2004). This shunt provides a constant supply of C to heterotrophic microorganisms in a dilute (40 μM C average) and highly viscous ocean, where the foraging distances are large and contain only a limiting nutrient supply to the microbial world (Fenchel, 1984; Jumars, 1993). Polymer gel networks are critical to providing a natural 3D matrix scaffolding capable of holding a heterotrophic microbial community together (Moon et al., 2007; Orellana et al., 2000), particularly in the deep ocean where non-­ sinking particles support microbial life (Baltar et al., 2010; Herndl and Reinthaler, 2013). Indeed, the presence of gels in the deep ocean may explain the discrepancy between bacterial C demand by the deep-water heterotrophic microbial community and the particulate organic C (POC) supply from the surface

461

ocean (Baltar et al., 2010; Herndl and Reinthaler, 2013). Polymer gels contain a rich nutritional microenvironment (polysaccharides, lipids, proteins, and nucleic acid chains; (Chin et al., 1998; Ding et al., 2008; Orellana et al., 2007), providing an optimal microenvironment of biodegradable substances—that is, a “hot spot” (Azam, 1998; Azam and Malfatti, 2007; Grossart et al., 2007; Malfatti and Azam, 2009) where heterotrophic bacteria/Archaea can interact and grow (Figure 9.2; Gram et al., 2002; Hmelo and Van Mooy, 2010; Malfatti and Azam, 2009). Bacterial growth is four to seven times faster when grown on gels than on free unassembled polymers (Orellana et al., 2000). Therefore, marine polymer gels form a fundamental structure and bacterial microenvironment (Malfatti and Azam, 2009). What is the relation between gel networks, lability, age, and the size of the DOC polymers in seawater? Based on turnover time, five fractions of DOC have been conceptualized (Carlson, 2002; Hansell, 2013), with four relevant here: (a) a highly labile pool of young polymeric material with a fast turnover lasting from hours to a few days; (b) a semi-labile fraction with turnover of months to a few years; (c) a semi-­ refractory fraction with lifetimes of a decade, and (d) a refractory, nonbiodegradable fraction with a turnover lasting thousands of years, exceeding the timescale of deep water circulation (an average of about 5000 years; see Chapter 6 and Bauer (2002)). These fractions overlap in distribution in the upper water column, with the refractory fraction being the most abundant and, chemically, the least understood (Hansell, 2013; Hansell and Carlson, 2013). The bioreactivity of marine polymers depends on many factors, including (a) the nature of the biopolymers (C:N ratio, amino acid composition) and their size (Amon and Benner, 1996), (b) the abundance and taxonomic composition of the colonizing microbial community, bearing the genetic capacity toward attachment to particles (Bauer, 2006; Carlson et al., 2004; Cottrell and Kirchman, 2000; DeLong et al., 2006), and (c) the functional genomic fingerprint

462

9.  Marine Microgels

of the bacterial population colonizing and degrading the gels and/or particles (Fuhrman, 2009; McCarren et al., 2010; Moran et al., 2007; Palenik et al., 2006). LMW DOC (<1000 Da) exhibits the lowest bioreactivity, while HMW DOC (>1000 Da but <0.2 μm) and colloids support the bulk of the marine heterotrophic microbial production, thus LMW DOC accumulates (Amon and Benner, 1994, 1996; Simon et al., 2002; Verdugo, 2012). These LMW short-chain chemical moieties are too large to be incorporated into the bacterial cell (>600 Da) (Weiss et al., 1991) but too short to assemble into stable microgels and so remain dispersed in solution as part of the refractory DOC. Additionally, we now have a better understanding of bacterial ecophysiological strategies and bacterial interactions among themselves, and other taxa (CuadradoSilva et al., 2013; Dobretsov et al., 2009; Gram et al., 2002; Hmelo and Van Mooy, 2010; Orellana et al., 2013), as well as of the structure of the organic matter continuum network (Malfatti and Azam, 2009). These fundamental characteristics and the interplay between the DOC field and bacteria imply that microbial oxidation of DOC depends significantly on the structure of the organic matter field and the quaternary conformation of larger molecules and/or networks of smaller chains forming an organic matter continuum, rather than on smaller dilute single free monomers or oligomer chains (DOC ~40-70 μM) (Azam, 1998; Malfatti and Azam, 2009; Orellana et al., 2000; Simon et al., 2002; Verdugo, 2012; Verdugo et al., 2004). Polymer gel networks, as hot spots, likely play a fundamental role in influencing the biogeochemical fate of C in the oceans, with consequences ranging from bacterial interactions in the microbial loop, especially in the deep ocean, to climate change (Aristegui et al., 2009; Azam, 1998; Baltar et al., 2010; Herndl and Reinthaler, 2013; Verdugo, 2012). When microbial heterotrophic assemblages colonize these 3D gel networks, they hydrolyze and crack the long-chain polymers with their ectoenzymes (Arnosti,

2011); this ongoing process affects the assembly/dispersion equilibrium of the marine gels. Physical fragmentation of biopolymers by UV photolysis (see Chapter 8; Mopper and Kieber, 2002; Orellana and Verdugo, 2003) or by bacterial enzymatic degradation (Benner and Kaiser, 2011) produce short-chain monomers and oligomers that do not interact to assemble into stable networks, and therefore they accumulate in the ocean. Thus, the natural process of spontaneous assembly of DOC polymers is disrupted and inhibited (Figure 9.4; Orellana and Verdugo, 2003; Orellana et al., 2000). Biological degradation will also disperse assembled microgels, driving short chain molecules to diffuse into a dilute DOC environment (Orellana and Verdugo, 2003). The molecular architecture of bacterial porins confirms that bacteria/Archaea can utilize only biopolymers <600 Da (Weiss et al., 1991), with some exceptions (Payne and Smith, 1994). Therefore, free polymer chains between 600 and 1000 Da are too large to pass through the microbial membrane porin channels for bacterial assimilation and metabolization and too short to assemble into stable networks that bacteria can colonize and degrade (Azam and Malfatti, 2007). Although we have no clear consensus on why old refractory DOC accumulates in the world oceans (Hansell, 2013; Jiao et al., 2010), soft matter polymer theory can explain the mechanisms by which short-chain polymers (<1000 Da) accumulate in the world oceans. Irrespective of their chemistry, monomers and oligomers remain dispersed (unassembled) in seawater due to their inability to form stable gels, since the probability of assembly of polymer gels depend on the square of the polymer length. Therefore, short-chain oligomers and probably complex molecules cycle slowly in the world oceans (Aluwihare et al., 1997; Bauer et al., 1992; Benner and Kaiser, 2011; Chin et al., 1998); unstable colloidal-size gels (100 and 200 nm) too small for microorganisms to colonize will have a similar character (Koike et al., 1990; Wells and Goldberg, 1992). The n ­ anocolloids accumulate



III  Structure, Properties, and Dynamics of Marine Polymer Gels

in the oceans, forming part of the refractory DOC (Benner, 2002). This refractory fraction also may include polymers that have undergone volume phase transition (see below) and/ or are chemically complex (Vandenbroucke and Largeau, 2007). 4  Macroscopic Polymer Gels and TEP Marine particles and their dynamics have long interested oceanographers (Paerl, 1973; Riley, 1963). Attention has grown as studies have brought new knowledge about colloid-sized particles—and their role in trace metal scavenging (Koike et al., 1990; Wells and Goldberg, 1992, 1993, 1994), reviewed by Wells (2002); marine snow and macroscopic particles such as TEP and gelatinous macro-aggregates—as well as their role in the biological pump (Alldredge et al., 1993; Burd and Jackson, 2009; Jackson and Burd, 1998; Jackson, 1990; Passow, 2002b). Today, scientific literature contains >1000 papers about TEP (Google Scholar). TEP (>0.7-500 μm particles) have been studied in vitro and in situ, from the Atlantic (Engel, 2004; Martin et al., 2011b) to the Pacific (Alldredge et al., 1993) to the Indian Oceans (Kumar et al., 1998), from tropical to temperate to polar (Hong et al., 1997) and from the sea surface microlayer (SML) to the deep ocean (Wurl et al., 2009). Abundant TEP also exist in freshwater lakes (Chateauvert et al., 2012; de Vicente et al., 2010; Pace et al., 2012). Marine particles have been investigated in the context of polymer physics (Chin et al., 1998), bringing new insights and understanding about their dynamics. Macroscopic gels assemble from large polymer chains held together by metal ions (Ca2+ (Chin et al., 1998; Ding et al., 2007; Orellana et al., 2007; Orellana and Verdugo, 2003; Verdugo, 2012; Verdugo and Santschi, 2010), hydrophobic polymers (Ding et al., 2008), and/or hydrogen bonds (Radić et al., 2011). TEP commonly form the matrix for the aggregation of cell debris and detritus and small organisms, arising mainly on the demise of phytoplankton blooms (Alldredge and Jackson, 1995; Logan

463

et al., 1995; Passow, 2002b). TEP contain a subgroup of all polymers encountered in the ocean (only acidic polysaccharides) and of polymers that assemble into marine gels (Verdugo, 2012; Verdugo and Santschi, 2010). TEP are operationally defined as acidic polysaccharides—­ especially sulfate ester groups (Mopper et al., 1995; Zhou et al., 1998b), carboxyl, sulfate, and phosphate-containing particles (Hung et al., 2001, 2003; Passow, 2002a, b) that can be visualized when stained with Alcian blue (a cationic copper phthalocyanine dye that complexes carboxyl (COO−) and half-ester sulfate (OSO reactive groups of acidic polysaccharides) at pH 2.5). Stained exopolymer particles can be measured in semiquantitative assays, either microscopically or colorimetrically, using xanthan gum equivalents (μg L−1), a proxy for marine polymers for calibration purposes, which makes the quantitative expression of these particles difficult to associate with measures of C export in the ocean and are rarely measured (Engel and Passow, 2001; Mari, 1999). Furthermore, as the C content of marine polymers per Alcian Blue binding site varies, the conversion between xanthan gum equivalent and C also varies; it needs to be ascertained for each analytical case (Engel and Passow, 2001; Mari, 1999; Mari et al., 2001). TEP-forming acidic polysaccharides are secreted by phytoplankton and bacteria, especially diatoms (Alldredge and Gotschalk, 1989; Mopper et al., 1995); however, phytoplankton groups such as dinoflagellates, cyanobacteria, and prymnesiophytes can produce large amounts of TEP-forming polymers as well (Alldredge et al., 1998; Hung et al., 2003; Passow, 2002b). TEP can form fast, in time frames of hours (Wurl et al., 2011), and can be exported swiftly into the water column and sediments, reaching average rates equal to 20 g C m2 day−1 (dry weight) (Logan et al., 1995) or 10-100 mg xanthum gum eq. m−2 day−1 (Passow et al., 2001). Thus, while TEP might have an important role in the biological pump, it suffers from its semiquantitative nature and the difficulty of transforming xanthan gum

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9.  Marine Microgels

e­ quivalent into actual C concentrations (Alkire et al., 2012; Martin et al., 2011b). TEP have been described as gels because of their gelatinous nature (Wurl and Holmes, 2008). While TEP might be gels, they have not been demonstrated to have the emergent physicochemical characteristics of polymer gel networks (spontaneous assembly, volume phase transition). Furthermore, the processes of aggregation of TEP are based on coagulation theory, collision, and stickiness (the probability of two particles adhering once they collide; Engel et al., 2004); absent is the mechanistic understanding of the intermolecular association, energies, and assembly processes of the polymers that form them (Verdugo, 2012). Molecular dynamic simulations are today used to model polymer aggregations, ranging from pure proteins to lipids to carbohydrates (Poon and Andelman, 2006). This approach could be used in the future to accurately simulate marine polymer dynamics and aggregation using the distributions of bonds, angles, and dihedrals (Lee et al., 2009). While these simulations have not been used in marine biogeochemistry, they might advance understanding of C dynamics, trace metal scavenging, and export into the deep ocean.

IV  PHASE TRANSITION All polymer gels have a remarkable feature: they undergo a reversible, mechanical deformation that does not affect the structure of the network. Polymer gels can undergo volume phase transition, from a swollen, hydrated phase to a condensed and compact phase. The gel volume can change several hundred times in response to infinitesimal changes and shifts in environmental conditions; such changes have been observed universally in synthetic and natural polymer gels (Pollack, 2001; Skubatz et al., 2013; Tanaka, 1992; Tanaka et al., 1980). Changes in volume occur in association with the internal dielectric properties of the gel polymer matrix in response

to environmental stimuli, on the nature of interactions between polymers and on the environment (Tanaka et al., 1980), such as temperature and UV light (Mamada et al., 1990); trace metals and pollutants (Jadhav et al., 2010; Rosen, 1993); and probably many other unknown chemicals. Volume phase transition in marine gels can be induced by changes in temperature (Verdugo, 2012; Verdugo and Santschi, 2010); pH (Chin et al., 1998; Orellana et al., 2011b); climate-relevant substances, such as dimethyl sulfide (DMS) and its precursor dimethylsulfoniopropionate (DMSP) (Orellana et al., 2011b); and polycations, such as polyamines (Nishibori et al., 2003). External stimuli induce marine microgel volume phase transitions (swelling and condensation) and the further collapse of the microgels by expulsion of water into a dense, compact polymeric network, increasing its specific weight and thus probably its sedimentation rate into the deep ocean; however, the settling rates of gels under these conditions have not been measured (Chin et al., 1998; Orellana and Hansell, 2012; Orellana et al., 2011b; Verdugo and Santschi, 2010). During the process of phase transition, molecules such as the photosynthetic protein RuBisCO (ribulose-1,5-bisphosphate carboxylase/oxygenase), DMSP, and DMS can be entrapped and packaged or caged at very high concentrations within the gel (Fernandez et al., 1991; Orellana and Hansell, 2012; Orellana et al., 2011a; Verdugo et al., 1995). These gels could then be carried to the deep ocean. In the Equatorial and North Pacific, for example, microgels containing immunologically recognizable RuBisCO were found at 3000 m (see below, Orellana and Hansell, 2012). The first demonstration of phase transition in marine gels (Chin et al., 1998) found marine microgels to collapse at pH 4.5; these gels exhibited a 20-fold volume collapse, from 5 μm to 250 nm. This pH corresponds with the pKa of carboxylic acid groups, consistent with measurements showing that one out of every six C residues in DOC corresponds to a carboxylic group (see



IV  Phase Transition

Chapter 2; Benner et al., 1992; Hertkorn et al., 2006; Lechtenfeld et al., 2014). Sulfuric acid (H2SO4), an oxidation product of the climate-­ relevant chemical DMS, also induces reversible volume phase transitions in microgels found at high latitudes in the high Arctic (Orellana et al., 2011b). In this case, the steep volume transition from the hydrated to the condensed phase took place as the pH decreased from 8 to 6. This pH transition inflection point is higher than observed (pH 4.5) for marine polymer gels from other latitudes (Chin et al., 1998), perhaps reflecting differences in polymer composition. The low transition pH measured by Chin et al. (1998) was used as an experimental demonstration that marine gels do indeed undergo discontinuous volume changes; but this value is not yet relevant in today’s changing and acidifying ocean (pH 8.05), even in corrosive upwelling areas (pH 7.6; Feely et al. 2008, 2009). However, H2SO4 is germane to nanometer-sized atmospheric particles (Leck and Bigg, 2005a). Furthermore, recent research has demonstrated that microgels containing and entrapping RuBisCO in the Pacific Ocean (Orellana and Hansell, 2012) undergo a steep volume condensation and phase transition in vitro. These gels collapse from a swollen network at pH 8 to a nonporous, tight network at pH 7, where they reach a size of at least six times smaller than that at pH 8. The volume phase transition of the microgels containing RuBisCO resulted in nonporous polymeric networks analogous to a stone-like particle smaller than 300 nm, probably preventing further bacterial enzymatic degradation by inhibiting diffusion of the enzymes into the collapsed network (Verdugo, 2012). One may expect that RuBisCO, as a soluble enzyme within algal chloroplasts (pH 8), would be biodegraded rather quickly by microzooplankton and/or zooplankton in the water column. But the pH inside the vacuoles of protozoa and in zooplankton guts can decrease abruptly with the production of degrading enzymes (Laybourn-Parry, 1984; Pond et al., 1995), inducing physicochemical alterations and

465

a microgel volume phase transition from swollen and hydrated to condensed and collapsed at pH 7. While these networks should swell once back in seawater, entrapped and caged in a gel, RuBisCO would be protected from microbial degradation (Verdugo et al., 1995), explaining the presence of algal RuBisCO in the deep Pacific Ocean (Orellana and Hansell, 2012). The combined effect of ocean pressure and temperature on marine proteins is not known. The process of phase transition can have important biogeochemical implications. Apparently, it prevents the degradation of proteins such as RuBisCO and probably other proteins present in the DOM pool by inhibiting microbial degradation caused by decreasing permeability of the collapsed network by exoenzymes (Orellana and Hansell, 2012; Verdugo, 2012). Phase transition could also explain how autotrophic biomolecules, specifically proteins, escape biodegradation in the water column, why the dissolved organic nitrogen pool, which exists mainly as amides in the interior of the ocean, resists decay, and why marine proteins are preserved in the deep ocean (Aluwihare et al., 2005; McCarthy et al., 2004; Orellana and Hansell, 2012); however, this protective mechanism needs to be demonstrated in situ. Volume condensation, followed by volume collapse of these polymer microgel networks can also be induced by micromolar levels of DMS and its biogenic precursor DMSP (Figure 9.7). The concentrations of DMS and DMSP inducing microgel phase transitions are consistent with extracellular and intracellular concentrations of these climate-relevant chemical compounds produced by polar phytoplankton and ice algae (Matrai and Vernet, 1997), as well as by phytoplankton from the Sargasso (Gabric et al., 2009) and the tropical Pacific (Bates and Quinn, 1997). The results showing microgels undergoing phase transition with DMS and DMSP are also analogous to the finding that high concentrations of DMSP and DMS are stored in condensed state in the acidic secretory vesicles of

466

9.  Marine Microgels 600 Number Volume

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FIGURE 9.7  Microgel volume phase transition. (a) pH. Marine polymer gels undergo fast, reversible volume phase transition (<1 min) from a swollen or hydrated phase to a condensed and collapsed phase by changing the pH of the seawater with H2SO4. The sizes of the polymer gels were monitored by confocal microscopy and the number of gels by flow cytometry. The swelling/condensation transition is reversible and has a steep sigmoidal change in the volume of the gels. Each datum corresponds to the average and standard deviation of three samples. (b) DMS and DMSP. Marine polymer gels can undergo fast, reversible change from a swollen/hydrated phase to a condensed phase as a function of DMS and DMSP concentrations; data expressed as the ratio between initial (Vi) and final (Vf) microgel volume, before and after addition of the inducing compound, respectively, and measured with confocal microscopy (Orellana et al., 2011b).

the Prymnesiophyceae microalga Phaeocystis in condensed state, within the polyanionic gel matrix (Orellana et al., 2011a). The secretory vesicles are stimulated by blue light (and probably other unknown stimuli) to release their concentrated content into the environment by the process of exocytosis (Chin et al., 2004). Secretion of polymer gels by this process is accompanied by elevated DMS and DMSP concentrations, suggesting that these substances are released with the polymer gel matrix into the water column (Matrai and Vernet, 1997) as well as into the clouds (Orellana et al., 2011b). Phase transition of freshwater gels has also been demonstrated at a critical point between

pH 6.5 and 7, with serious consequences for the optical landscape of the organic matter field (Pace et al., 2012). At low pH, as these authors demonstrated, DOC polymers and colloids are condensed and compact. This volume condensation affects the optical characteristics of colored DOM (CDOM) by limiting the exposure of chromophores to light; conversely, at higher pH, polymers and colloids expand, exposing chromophores to light. This experimental dynamic change in volume of the freshwater gels resulted in changes of light absorption and photobleaching, which affects water transparency and ultimately may strongly influence C cycling, including the balance of autotrophy



V  Marine Gels in the Atmosphere and Their Relevance for Cloud Formation

and heterotrophy in freshwater ecosystems (Pace et al., 2012). However, the effects of pH on the structure of the CDOM are different in marine waters. Marine waters do not show big changes in pH except in microenvironments or diurnal changes in pH related to photosynthetic processes (Cornwall et al., 2013; Schmalz and Swanson, 1969). Volume phase transitions of CDOM may also be induced by changes in salinity gradients along estuaries, as indicated by discontinuous transitions in maximal fluorescence emission by CDOM when small changes in salinity (33-35) occurred in samples taken from Puget Sound, the Orinoco River plume, and the west Florida shelf (Del Castillo et al., 2000). Additionally, volume phase transitions driven by hydrophobic interactions can be induced by temperature changes, with critical transition points at 5 °C and from 24 to 30 °C (Verdugo, 2012). Finally, volume phase changes may also be implicated in the production of biogeocondensates in sediments, where large changes in in situ pH and other chemical parameters are known (Fenchel et al., 2012), preventing microbial exoenzymatic degradation of kerogen and algaeans (Vandenbroucke and Largeau, 2007).

V  MARINE GELS IN THE ATMOSPHERE AND THEIR RELEVANCE FOR CLOUD FORMATION Clouds remain a weakness in our understanding of the climate system and consequently in climate modeling (IPCC, 2007). Clouds form when water vapor condenses. But, water vapor needs something to condense on—tiny airborne aerosol particles known as CCN. Typically, CCN fall within the submicron size fraction, about 100 nm in diameter. Depending on their properties and heights, clouds can either warm their surfaces by triggering a localized greenhouse effect or cool them by reflecting solar radiation. If CCN are scarce, the resulting clouds will c­ontain fewer and larger d ­ roplets.

467

Such clouds will reflect little sunlight to space while blocking the escape of heat from Earth’s surface, causing it to warm. However, if CCN are plentiful, many fine droplets form; the resultant clouds are better reflectors, thus cooling the surface below. Over remote marine areas such as in the high Arctic (>87oN), anthropogenic particles are virtually absent. Instead, biological sources of particles may dominate (Bigg and Leck, 2001a,b; Leck and Persson, 1996; Leck and Bigg, 2005a,b; Leck et al., 2002, 2013; Orellana et al., 2011b). This “clean” air, with few CCN, makes the low-level stratocumulus clouds optically thin, with fewer but larger droplets. But, if a warming climate spurs the activity of microbiota, organic sources of CCN might become more prominent and lead to increasingly r­ adiation-reflective clouds. Over the last 15 years, articles have emphasized the presence and enrichment of organic matter particles of submicron sizes in airborne aerosols and cloud water (Bigg and Leck, 2001, 2008; Leck and Bigg, 2008 and Karl et al., 2013; Duce and Hoffman, 1976; Facchini et al., 2008; Gaston et al., 2011; Keene et al., 2007; Leck and Bigg, 2005a,b, 2010; Middlebrook et al., 1998; O’Dowd et al., 2004; Russell et al., 2010; Yoon et al., 2007). The detection of organic substances—­specifically of exopolymer like particles—in the atmosphere was first discovered by Bigg and Leck (Bigg and Leck, 2001a,b; Leck and Bigg, 2008; Bigg and Leck, 2008; Leck and Bigg, 2005a; Leck et al., 2002). These authors recognized that these particles could bear the physicochemical characteristics of marine gels. This followed from their studies of a possible link between cloud formation and polymer gels in the SML (<100 μm thick at the air-sea interface) in the high Arctic sea ice (Bigg et al., 2004).

A  Is the “Gel Theory of Marine CCN” Coupled to the Sulfur Cycle? Charlson et al. (1987) reviewed existing evidence that implicated DMS (produced by the microbial food webs) in the production of CCN over remote marine areas. This provocative CLAW hypothesis

9.  Marine Microgels

FIGURE 9.8  A polymer gel microcolloid collected in the air over the Arctic pack ice at 89° N. Adapted from Leck and Bigg (2005a).

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(so named informally after the paper’s authors Charlson, Lovelock, Andreae, and Warren) stated that, in the marine domain, DMS emissions and their subsequent oxidation products—methane sulfonic acid, sulfur dioxide, and sulfuric acid— trigger cloud formation, which cools the ocean surface. This cooling would, in turn, affect further emissions of DMS by changing the speciation/ abundance of marine phytoplankton, establishing a feedback loop. Observations in the early 1990s from the Arctic did indeed show that the intermediate oxidation products provided most of the mass for the CCN-sized particles observed over pack ice (Leck and Persson, 1996). The source of most of the DMS, though, was found at the fringe of the central Arctic Ocean, at the hospitable edges of the pack ice. At that time, this distribution suggested that winds carried DMS-rich air toward the North Pole, and oxidation of this DMS created extremely small sulfuric acid particles. Theoretically, these particles would then grow slowly by further condensation of the acids until they were large enough to serve as CCN. Surprisingly, it turns out that sulfuric acid had nothing to do with the small precursors of CCN. Instead, observations from the Arctic in the mid1990s showed that these small precursors were mostly particles resembling viruses and nanogels and microgels that were accompanied by other larger particles, such as bacteria and fragments of diatoms (Figure 9.8; Bigg and Leck, 2001a,b; Leck and Bigg, 1999; Leck and Bigg, 2005a, 2010; Leck et al., 2002). Subsequently, Bigg et al. (2004) detected large numbers (106-1014 mL−1) of similar particles within the thin surface film at the water-air interface between ice floes (Figure 9.9). We know that polymer gel networks assemble preferentially at the water-air interface (Verdugo et al., 2004). The SML has long been known as a source of gels (Sieburth, 1983), but the connection between the airborne particles and the SML was found much more recently. Airborne microgels may have the chemical surfactant properties necessary to act as nuclei for clouds (CCN), but to behave as effective CCN,

Relative frequency (%)

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FIGURE 9.9  Red curve. With standard error bars: ­median particle number size distribution of five SML ­ samples ­expressed as relative frequency of occurrence (right y-axis). Black curve, relative particle number size distribution of the individual components of the particles presented in the median size distribution (left y-axis). Black dotted curve not described. Inset, upper right. Microscopic image of particles from samples, corresponding to the red curve. Inset, upper left. Microscopic detail of an aggregated particle typical of those sized in the red curve. Modified from Bigg et al. (2004).



V  Marine Gels in the Atmosphere and Their Relevance for Cloud Formation

they must reach a critical size and meet other physicochemical properties and energy constraints of the system. Leck and Bigg (1999, 2010) and Karl et al. (2013) speculated that the primary marine gel would disintegrate under some circumstances, generating smaller particles, probably due to UV radiation cleavage (Orellana and Verdugo, 2003). However, it is not clear whether the polymeric material reached smaller sizes due to breakage caused by UV radiation, by reversible volume phase transition, or by a combination of both, as has been described for marine microgels (Chin et al., 1998; Orellana et al., 2011b; Orellana and Verdugo, 2003); thus, this remains an open question. Furthermore, the gels could also provide sites for condensation of the oxidation products of DMS. In 2005, when Leck and Bigg tested predominantly airborne sulfate particles for the presence of microgels, they detected marine microgel material in half or more of their samples coated with sulfuric acid. A specific fluorescently labeled antibody probe developed against in situ seawater and SML biopolymers confirmed for the first time that the particles found in the atmosphere (aerosol/fog/cloud) originated in the surface sea water, including the subsurface and SML (Orellana et al., 2011b). The particles were released by sea-ice algae, phytoplankton, and bacteria, and behaved as nanogels and microgels (Figure 9.10). The gel networks were held together by random entanglements and Ca2+ ionic bonds (Figure 9.3), as well as by hydrophobic moieties (Figure 9.5). The gels comprised as much as 50% of the total organic C in surface waters and the SLM, and they assembled faster than previously observed, probably due to the presence of hydrophobic moieties enhancing polymer assembly (Figure 9.4; Ding et al., 2008). The gels also underwent volume phase transitions induced by DMSP as well as DMS, another indication that those particles displayed the physicochemical characteristics of gels (Figure 9.7). Gels were abundant in the subsurface seawater and enriched in the SML. They also correlated with enrichment of proteins containing hydrophobic amino acids and DMSP in the SML.

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FIGURE 9.10  Cloud water: polymer gels immune-­ labeled with a fluorescently labeled antibody developed against seawater biopolymers (Orellana et al., 2011b).

The co-occurrence of atmospheric organic material and biologically active marine waters has, since the mid-1990s, been confirmed for the high Arctic waters, but it has also been documented for temperate waters (Facchini et al., 2008; Leck and Bigg, 2008, 2010; Russell et al., 2010). Observations from the Arctic called into question the key role given to DMS in the CLAW hypothesis (Leck and Bigg, 2007). In the emerging picture of the Arctic atmosphere, DMS concentrations will determine the mass of the particles by producing material for their growth. But, it is the number of airborne gels that will primarily influence the number of CCN and the resulting optical properties of the cloud droplets. Indeed, research during the past two decades—reviewed by Quinn and Bates (2011)—does not corroborate the CLAW hypothesis for other regions either.

B  The Effect of Gels on Bubble Properties and Bursting Leck and Bigg (Leck et al., 2004; Leck and Bigg, 2005b and Bigg and Leck, 2008; Leck and Bigg, 1999; Leck et al., 2002) had hypothesized that the source of gels found in clouds was the open

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9.  Marine Microgels

water between ice floes and that those particles would be transferred to the atmosphere by the bursting of air bubbles at the air-sea interface. Transport of aerosols by bubble bursting under experimental conditions is well known (e.g., Aller et al., 2005; Fuentes et al., 2010a; Fuentes et al., 2010b; Fuentes et al., 2011; Keene et al., 2007; Kuznetsova et al., 2005). However, experimental setups and different conditions show enormous variability as to aerosol size and concentration. Factors that affect the variability include the presence of surfactants, salinity, viscosity, Langmuir circulation, turbulence, wave breaking, etc. (Fuentes et al., 2010a). Bubbles usually result from entrainment of air induced by wind stress at the air-water i­ nterface (Blanchard, 1971; Blanchard and Syzdek, 1988); bubble bursting produces primary aerosol particles in CCN sizes. Over the summer pack ice, near-surface wind speeds are typically low (<6 m s−1), and the extent of open water in the pack ice leads is usually modest (10-30%), shortening fetches and limiting the generation of waves. A recent study confirmed, in spite of the low winds, both the presence and the temporal variability of a population of bubbles within the open leads; proposed was a non-wave bubble source mechanism; the loss of heat to the atmosphere produced temperature differences at the surface layer and bulk water creating convective mixing and driving local fluctuations in gas saturation, that subsequently generated both film and jet droplets (Norris et al., 2011). Possible nonwind-related sources of bubbles include releases of bubbles trapped in melting sea ice, as well as those expelled by freezing water, or transported to the surface by increased turbulence caused by super-cooling conditions (Grammatika and Zimmerman, 2001). Further sources of bubbles include photosynthesis and respiration of phytoplankton (Johnson and Wangersky, 1987), and falling raindrops on the ocean surface (Lewis and Schwartz, 2004). Bubbles scavenge DOM to the bubble film (Zhou et al., 1998b), particularly demonstrating

the role of surface-active polysaccharides in the formation of large TEP by bubble adsorption in seawater. However, during bubble bursting over marine areas, bubbles scavenge not only debris and HMW soluble organic surface-active compounds but also sea salt, as they rise through the water prior to their injection into the atmosphere (Blanchard and Syzdek, 1988). It has generally been assumed that particles derived from bubble bursting would be composed of sea salt only, and would thus contribute a significant fraction of the CCN population (O’Dowd et al., 1999). However, transmission electron microscopy of individual particles (Bigg, 1980; Bigg and Leck, 2001, 2008; Gras and Ayers, 1983; Leck and Bigg, 2005a,b; Leck et al., 2002; Pósfai et al., 2003) over the pristine perennial arctic ice, and at remote marine locations at lower latitudes, have failed to find evidence of sea salt particles <200 nm in diameter. To explain this, Bigg and Leck (2008) proposed a mechanism for transporting polymer-­microgel-rich organic material from the bulk seawater into the open lead SML. They suggested that the highly surface-active polymer gels could concentrate on the surface of rising bubbles and then aggregate by interpenetration and polymer tangling. Consequently, rising bubbles can selectively carry polymer gels, as well as embedded solid particles such as bacteria and phytoplankton and their detritus, to the SML. Before bursting, bubbles rest in the microlayer; therefore, they likely have a film envelope composed largely of gels, and embedded particulate matter that may become points of weakness as the water drains from between their envelope. Following the burst, the film drops fragments, containing surfactant material, salt-free w ­ ater, and any particle attached to the fragments. These suggestions are consistent with the fact that gels assemble preferentially at the microlayer interface (Verdugo et al., 2004) as well as on bubble films (Gao et al., 2012). An alternative process for expelling polymer gels from the ocean surface involves charge repulsion by the negatively charged surface of



V  Marine Gels in the Atmosphere and Their Relevance for Cloud Formation

the Earth, which acts as a spherical capacitor. The Earth has a net negative charge of about a million coulombs, and an equal positive charge resides in the atmosphere (Feynman et al., 1964). No observational measurement to explain this path of the aerosols in the atmosphere has been done.

C  Is the Gel Theory for the Origin of Marine CCN Consistent with Primary Marine Aerosol Observations? The Arctic studies of primary organic polymer gel aggregates derived from the SML between sea ice leads, performed over the last two decades, have been expanded to other marine areas (Facchini et al., 2008; Fuentes et al., 2010b; Leck and Bigg, 2008; Russell et al., 2010). These studies show strong similarities in the morphological and chemical characteristics of the particles both previously and recently described (Leck and Bigg, 2005b, 2007, 2008; Leck et al., 2013). Most of the studies have focused on size-­ segregated chemical characterization of marine aerosol particles (Cavalli, 2004; Keene et al., 2007; O’Dowd et al., 2004; Rinaldi et al., 2010; Russell et al., 2010; Sciare et al., 2009). They all have suggested marine DOM at the ocean surface as the source for these particles. However, they have not looked at gel’s physical-chemical characteristics and the emergent properties (assembly, phase transition) of this sort of particulate matter and thus have not confirmed that they are marine gels. Recently, Martin et al. (2011a) suggested that internally mixed amphiphilic biopolymers and gels do not have the ability to activate as CCN, instead CCN were dominated by the sulfate fraction of the particles, which could be trapped within the polyanionic matrix of the gel (Leck and Bigg, 2005a; Orellana et al., 2011a). However, we know that marine gels have the important property of being solvated in water, highly surface active and also of undergoing ­reversible volume phase t­ ransitions. Marine gels

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can reversibly change their morphology from a ­swollen, hydrated phase to a condensed, collapsed phase. The volume phase transition may, for instance, be caused by changes of pH, DMS, and DMSP (Orellana et al., 2011b). Moreover, Arctic gels were shown to consist of hydrophilic and hydrophobic segments (Orellana et al. 2011b) in agreement with their chemical behavior. This behavior was recently confirmed by molecular dynamic modeling by (Xin et al., 2013). Hence, the colloidal gel shows only a partial wetting character below 100 percent relative humidity (RH) thus showing only weak hygroscopic growth but, at the same time, high CCN activation efficiency is shown at RH above 100 percent. (Orellana et al., 2011b). Experimental data reported by Ovadnevaite et al. (2011) suggest a dichotomous behavior for the primary marine organic aerosol; water vapor does not uniformly undergo condensation, since only part of the surface exhibits strong hydrophilicity. Hence, the colloidal gel shows only a partial wetting character (<100% relative humidity), thus showing only weak hygroscopic growth but, at the same time, high CCN activation efficiency. The 3D structure of marine microgels, with their hydrophilicity or surface-active properties and only partial wetting character that result from the hydrophobic characteristics of the polymers, could explain such dichotomous behavior. Our understanding of gels and their emergent behavior as CCN is still in its infancy; however, the gel conceptual framework provides a predictive theory (Edwards, 1986; Tanaka, 1981, 1992) for understanding soft matter colloidal processes and controlling factors. While, we have no observations of gels as CCN outside the high Arctic, these observations suggest this might be a ­universal process.

Acknowledgments We thank the editors, D.A. Hansell and C.A. Carlson, for the invitation to prepare this chapter on marine microgels. The chapter benefited greatly from two anonymous reviewers for helpful comments, and the NSF Biological Oceanography

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Program, the Arctic Natural Sciences Program, the Polar Science Center (University of Washington), the Institute for Systems Biology, the Swedish Research Council, mentoring by Pedro Verdugo, and collaborations with Patricia Matrai, Keith Bigg, Robert Moritz and many students.

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