Biodegradation of Polymeric Mulch Films in Agricultural Soils

Perspective is published by the Biodegradation of American Chemical Society. 1155 Sixteenth Polymeric Mulch Films Street N.W., Washington, DC 20036 in Agricultural Soils: Publishedby byUniversity American of Subscriber access provided Chemical Society. Library Wollongong Copyright © American Chemical Society. However, no copyright

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MichaelDC Sander 20036

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Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b05208 • is published by the American Publication Date (Web): 30Chemical Jan 2019

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Page 1 ofEnvironmental 31 Science & Technology

Cpolymer

?

CCO + Cmicrobial biomass (+ Cremnant polymer) 2

keyACSsteps inPlus polymer film Paragon Environment biodegradation in soils

Environmental Science & Technology

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Biodegradation of Polymeric Mulch Films in Agricultural Soils:

3

Concepts, Knowledge Gaps, and Future Research Directions

4 5

MICHAEL SANDER*

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Institute of Biogeochemistry and Pollutant Dynamics

8

ETH Zurich

9

8092 Zurich, Switzerland

10 11 12

Submitted as perspective to Environmental Science & Technology

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*Corresponding author:

15

Michael Sander

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Email: [email protected]

17

Phone: +41 44 632 83 14

18 19

Number of pages: 29

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Number of figures: 4

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Number of tables: 0

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Number of words: 4086

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ACS Paragon Plus Environment

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Abstract

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The agricultural use of conventional, polyethylene-based mulch films leads to the

28

accumulation of remnant film pieces in agricultural soils with negative impacts for soil

29

productivity and ecology. A viable strategy to overcome this accumulation is to replace

30

conventional with biodegradable mulch films composed of polymers designed to be degraded

31

by soil microorganisms. However, understanding polymer biodegradation in soils remains a

32

significant challenge due to its dependence on polymer properties, soil characteristics and

33

prevailing environmental conditions. This perspective aims to advance our understanding of

34

the three fundamental steps underlying biodegradation of mulch films in agricultural soils:

35

colonization of the polymer film surfaces by soil microorganisms, depolymerization of the

36

polymer films by extracellular microbial hydrolases, and subsequent microbial assimilation

37

and utilization of the hydrolysis products for energy production and biomass formation. The

38

perspective synthesizes the current conceptual understanding of these steps and highlights

39

existing knowledge gaps. The discussion addresses future research and analytical

40

advancements required to overcome the knowledge gaps and to identify the key polymer

41

properties and soil characteristics governing mulch film biodegradation in agricultural soils.

42 43

Introduction

44

In order to secure food for the growing world population,1,2 modern agriculture heavily

45

relies on the use of plastic films in diverse applications, including soil mulching, greenhouses,

46

low and high tunnels, and silage.3-6 The global agricultural film market is predicted to reach an

47

annual volume of 7.5 million tons by 2021 with mulch films having a major share (> 40%).7-9

48

Conventional mulch films are composed of polyethylene (PE)3,6,10 and have thicknesses

49

between a few to several tens of µm. When applied to agricultural soils, these films raise crop


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yields by elevating soil temperatures, conserving soil moisture, controlling weed growth, and

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providing protection against severe weather and pests.6,8,11-13 Currently, China uses the most

52

PE mulch film, with an estimated 1.25 to 1.4 million tons of film applied annually,14,15 covering

53

approximately 20 million hectares or 12% of China’s farmland.14,16,17 This area equals about

54

five times the total size of Switzerland.

55

Recovery of PE mulch films from agricultural fields after use is often incomplete due to

56

PE film embrittlement and fragmentation caused by weathering, particularly when thin films

57

are used.4,16,18 Residual films enter and subsequently accumulate in agricultural soils because

58

PE is recalcitrant.19-21 Such accumulation is best documented for soils in the Xinjiang region

59

in northwest China, where intense mulch film application has led to residual PE film

60

concentrations in soils in the range of 120–350 kg residual film per ha and as high as 500 kg

61

residual film per ha.16,18,22 The accumulation of residual PE mulch films in agricultural soils

62

around the world has raised concerns because it decreases soil productivity by blocking water

63

infiltration, impeding soil gas exchange, constraining root growth, and altering soil microbial

64

community structures. 14,23 Besides impacting soil productivity, plastic pollution of soils is also

65

considered a general emerging threat to soil ecosystem health and function.24-30

66

A promising approach to overcome the accumulation of residual PE mulch films in soils

67

is to replace the conventional with biodegradable mulch films composed of polymers designed

68

to be degradable by soil microorganisms.31,32 As a promising technology to help overcome

69

plastic pollution of soils, biodegradable mulch films have also recently received increased

70

attention from regulators and scientists alike. In July 2018, a new EU standard was issued

71

specifying test methods and biodegradability criteria for biodegradable mulch films.33 Such

72

standards counteract commercialization of mulch films labeled as biodegradable without

73

scientific evidence in support of this claim (e.g., so-called ‘oxo-degradable’ PE films).19,34-38

74

Growing interest from the scientific community on the use of biodegradable mulch films is


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reflected in a number of recently-published reviews, policy communications, and viewpoints

76

that address standards and test systems for biodegradable mulch films, film properties and soil

77

characteristics that affect biodegradation, as well as potential effects of mulch film applications

78

on soil microbial ecology and overall soil function.9,12,13,39-43 A review that explicitly focuses

79

on the fundamental steps underlying biodegradation of mulch films in soils remained missing.

80

This perspective aims to advance a more holistic picture of three fundamental steps

81

underlying mulch film biodegradation in soils: The colonization of the polymer surfaces by

82

soil microorganisms (step 1), enzymatic depolymerization of the polymer by extracellular

83

hydrolases secreted by the colonizing microorganisms (step 2), and the microbial utilization of

84

the oligomeric and monomeric hydrolysis products that are released from the polymer (step 3).

85

To this end, the perspective summarizes the current conceptual understanding of each of the

86

three steps and highlights existing knowledge gaps, research needs, and associated analytical

87

challenges. Detailed knowledge of these steps is important not only to aid design and

88

mechanistically interpret experimental biodegradation studies but also to help develop

89

materials and application strategies that favor mulch film biodegradation in soils.

90

Biodegradable polymers and their use in mulch films

91

Biodegradable polymers are synthetic or natural organic polymers that degrade under the

92

active involvement of microorganisms.44,45 Biodegradation under aerobic conditions results in

93

the conversion of polymer carbon (𝐶𝑝𝑜𝑙𝑦𝑚𝑒𝑟) into carbon dioxide (𝐶𝐶𝑂2) and microbial biomass

94

(𝐶𝑏𝑖𝑜𝑚𝑎𝑠𝑠) (Eq. 1): 46,47

95

𝐶𝑝𝑜𝑙𝑦𝑚𝑒𝑟 + 𝑂2 →𝐶𝐶𝑂2 + 𝐶𝑏𝑖𝑜𝑚𝑎𝑠𝑠 ( + 𝐶𝑟𝑒𝑠𝑖𝑑𝑢𝑎𝑙 𝑝𝑜𝑙𝑦𝑚𝑒𝑟)

96

where 𝐶𝑟𝑒𝑠𝑖𝑑𝑢𝑎𝑙 𝑝𝑜𝑙𝑦𝑚𝑒𝑟 represents the carbon remaining in residual polymer as long as

97

Eq. 1

biodegradation is incomplete.

98

Biodegradability of a polymer depends on its physicochemical properties and not on the

99

provenance of its carbon (i.e., fossil-based vs. bio-based materials).40,45 Furthermore,


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biodegradability is a polymer trait that is specific to a given receiving environment (e.g.,

101

agricultural soils for mulch films).40,48-51 At the same time, polymer biodegradability varies

102

greatly between soils,52 demonstrating the importance of soil-specific characteristics in

103

polymer biodegradation.48

104

Figure 1a shows the chemical structures of both synthetic and natural polymers used in

105

commercial biodegradable mulch films.12,53,54 The synthetic polymers include the aliphatic

106

polyesters poly(butylene succinate), poly(butylene succinate-co-adipate) (PBSA), poly (-

107

caprolactone), and polylactic acid, as well as the aromatic-aliphatic co-polyester poly(butylene

108

adipate-co-terephthalate) (PBAT). Natural polymers include starch, cellulose and

109

polyhydroxyalkanoates. Most biodegradable mulch films are blends of two (or more) of the

110

listed polymers. A common chemical feature among the listed polymers is that their backbone

111

chains contain functional groups that are enzymatically hydrolyzable (i.e., a glycosidic bond

112

in starch and ester bonds in all other polymers; as shown for PBS and starch in Figure 1b).

113

Polyesters are widely used in biodegradable mulch films due to their desirable biodegradability

114

traits, comparatively low production costs and favorable thermal and mechanical

115

properties.55,56 Furthermore, altering the monomer composition of polyesters enables tuning of

116

degradability and performance traits for a particular application. For instance, increasing the

117

molar ratio of terephthalate to adipate increases the mechanical stability of PBAT but lowers

118

its biodegradability.57-59 This example highlights one of the most fundamental challenges in

119

developing biodegradable mulch films: balancing stability —needed for the films to fulfill their

120

function for the duration of the application to soils— and biodegradability —to ensure rapid

121

and extensive conversion to CO2 and microbial biomass once the films enter soils after use.

122


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123 124 125 126 127 128 129 130


Figure 1. a. Chemical structures of synthetic and natural polymers used in commercial biodegradable mulch films. The backbone chains of all polymers contain functional groups that can be enzymatically (and abiotically) hydrolyzed, a key step in polymer biodegradation. b. Depiction of the enzymatic hydrolysis of an ester bond in poly(butylene succinate) (PBS) (top) and an glycosidic bond in starch (bottom).

131

Steps involved in mulch film biodegradation in soils

132

This perspective addresses the complexity of mulch film biodegradation in soils by

133

separately examining the three fundamental steps that underlie this process (Figure 2). While

134

discussed separately, these steps are strongly interconnected and co-occur during mulch film

135

biodegradation. The discussion aims at identifying general concepts that broadly apply to

136

biodegradable mulch films irrespective of their exact polymeric composition. For this reason,

137

the generic term ‘polymer’ is used in the subsequent discussion when referring to synthetic and

138

natural polymers used in biodegradable mulch films. While pertinent, a discussion of the fate


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of mulch film additives (e.g., UV stabilizers, plasticizers, fillers, and synthesis catalysts) in

140

soils is beyond the scope of this perspective.

141

142 143 144 145 146 147 148 149

Figure 2. Three fundamental steps involved in polymer biodegradation in soils. Step 1: Microbial colonization of the polymer surface by film-degrading soil bacteria and fungi. Step 2: Enzymatic depolymerization of the polymer by extracellular hydrolases secreted by filmdegrading microorganisms colonizing the film surface. Step 3: Microbial uptake and utilization of monomers and short oligomers for energy production under release of CO2 and for biomass formation. Adapted from Zumstein et al., 2018.60

150

Step 1. Microbial colonization of polymer surfaces. The first step in the biodegradation

151

is the colonization of the polymer surface by degrading soil microorganisms that secrete

152

extracellular de-polymerases. Factors that facilitate colonization increase the total contact area

153

between microbial degraders and polymers and are thus expected to increase overall polymer

154

biodegradation rates.61

155

Conceptual understanding. Both fungi and unicellular organisms colonize the surfaces

156

of biodegradable polymers in soils, as shown by scanning electron microscopy (SEM)


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imaging.62-67 While microbial colonization is no proof for biodegradation —microbes also

158

colonize non-biodegradable polymers in the environment68-71— SEM images collected on

159

biodegradable polymers often show enhanced surface erosion around the colonizing

160

microorganisms,62-65 suggesting depolymerization by secreted hydrolases in proximity to the

161

colonizing microbes. Figure 3 shows exemplary SEM images visualizing colonization of

162

PBAT films by both fungi and unicellular organisms after six weeks of soil incubation.

163

A number of studies have characterized the microbial communities forming on the

164

surfaces of biodegradable polymers during soil incubations.62,72-75 Communities on the

165

polymer were enriched in specific fungi (often of the phylum Ascomyceta) and had a lower

166

fungal diversity as compared to the bulk soils.62,72,76 These findings likely indicate a

167

preferential colonization and proliferation by polymer-degrading fungi,72 which is supported

168

by the isolation of fungal degraders from the surfaces of biodegradable polymers incubated in

169

soils.63,64,77,78 Bacterial polymer degraders were also identified on the surfaces of biodegradable

170

polymers after soil incubation.74,79 Yet, the presence of biodegradable polymers in soils had

171

only small effects on the bacterial community composition and abundance in bulk soils.72,73,80

172

Taken together, microbial community analyses demonstrated colonization of polymers by both

173

fungal and bacterial degraders in soils.

174


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175 176 177 178 179 180 181 182 183

Figure 3. Selected scanning electron microscopy images of the surfaces of poly(butylene adipate-co-terephthalate) (PBAT) films that were incubated in an agricultural soil in the laboratory (six weeks; temperature of 25°C). The images serve to illustrate colonization of PBAT films by both fungi and unicellular organisms and to visualize hyphal growth along and into the polymer surfaces. Images in panels a and c were collected at different spots on the PBAT film surface. Panels b and d show magnifications of areas highlighted in panels a and c. Images were collected by Michael Zumstein.

184

Knowledge gaps and research needs. The spatiotemporal dynamics of fungal and

185

bacterial colonization have not been systematically investigated. SEM can be used to obtain

186

high-resolution images of microbial colonization —including potential co-localization patters

187

of bacteria and fungi— and associated (bio)erosion features on the film surfaces. SEM imaging

188

ought to be complemented by light and fluorescence microscopy approaches which have

189

potential to allow following microbial colonization dynamics in real time. Of particular interest

190

is fluorescence in situ hybridization (FISH) coupled to fluorescence microscopy69,81-83 because

191

this approach not only allows visualizing colonization but also provides information on the


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phylogenetic affiliation of colonizing microorganisms. FISH probes can be designed to screen

193

broadly for different phylogenetical affiliations of colonizing microorganisms. At the same

194

time, the use of FISH probes targeting specific fungal and bacterial degraders previously

195

identified in single culture or metagenomic studies would enable assessing their abundance on

196

polymer films in soils.

197

Complementary microscopy techniques can be used to systematically investigate the

198

effects of soil properties on microbial polymer colonization dynamics. Among these properties,

199

soil nutrient availability likely is of particular importance. Because the polymers used in

200

biodegradable mulch films are solely composed of C, O, and H (Figure 1), colonizers need to

201

acquire nitrogen (N) from the surrounding soil for growth. Therefore, colonizers may

202

experience N limitations in nutrient-poor soils and/or on the surfaces of rapidly depolymerizing

203

polymers that supply excess carbon relative to N to microorganisms. Compared to bacteria,

204

fungi may be less susceptible to N limitations. First, fungi have higher biomass C:N ratios of

205

approximately 15 as compared to 5 for bacteria, implying that fungi require comparatively

206

lower amounts of N for growth.84 Second, fungi can also use the cytoplastic current in their

207

hyphal network to direct N from nutrient rich microenvironments in soils to cells located on

208

the polymer surface.85 Limited N availability in specific soils may thus favor colonization by

209

fungi and possibly also heterotrophic nitrogen-fixing bacteria.86

210

Besides the stoichiometry considerations above, fungi have the additional advantage that

211

they can use directed hyphal growth to colonize film surfaces.63 Because hyphae can grow

212

across air gaps between soil particles and the film surface, fungi do not require direct contact

213

of soil with polymer films to initiate colonization. By contrast, unicellular bacteria likely

214

colonize the polymer surface primarily from soil-polymer contact points and through cell

215

division. The smooth and apolar film surfaces likely impair the formation of water films of

216

sufficient thickness and continuity required for bacterial motility as a means of


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colonization.58,87-89 However, it is conceivable that motile bacteria use the hyphal network on

218

the polymer surface for colonization (expressed in the ‘fungal highway’ concept).90,91 Based

219

on the above considerations, a testable hypothesis for future studies is that soil fungi (and

220

possibly also actinobacteria with filamentous growth92) outcompete unicellular bacteria in film

221

colonization.

222

Step 2: Enzymatic depolymerization of polymers. Microbial colonizers secrete

223

depolymerases to cleave the hydrolyzable bonds in the polymer structure, resulting in the

224

release of microbially-assimilable, low-molecular weight (i.e., < 600 Da)93 mono- and

225

oligomers (Figure 2). Abiotic hydrolysis of these bonds is typically much slower than

226

enzymatic hydrolysis under the temperature and pH conditions that prevail in soils.50,94,95

227

Enzymatic depolymerization is commonly considered the rate-limiting step in polymer

228

biodegradation in soils. This is supported by much faster microbial utilization of oligo- and

229

monomers when directly added to soils than of the corresponding polymers in the same

230

soils.60,96

231

Conceptual understanding. Enzymatic polyester hydrolysis has been extensively studied

232

because of its relevance to both polyester biodegradation in natural environments and polyester

233

recycling in engineered systems.97,98 To study this reaction, numerous analytical approaches

234

are available that quantify the release of hydrolysis products or protons into solution (one

235

proton is released per hydrolyzed ester bond at circumneutral pH),99-103 or increases in solution

236

fluorescence resulting from enzymatic co-hydrolysis of fluorogenic ester probes embedded

237

into the polyester.104 Enzymatic polyester hydrolysis has also been studied at the nanometer-

238

scale using atomic force microscopy105 and quartz-crystal microbalance measurements.106,107

239

Among several polymer properties controlling its enzymatic hydrolyzability,

240

crystallinity arguably is the most important.59,98,108-114 Polyesters used in biodegradable mulch

241

films are semi-crystalline and thus contain both amorphous and nm-sized crystalline


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phases.115,116 In the amorphous phases, polyester backbone chains are disordered and have a

243

higher mobility, favoring binding and hydrolytic cleavage by esterases. Contrastingly, the

244

polyester chains in crystalline phases are in highly-ordered arrangements with strong chain-

245

chain interactions. The resulting low mobility of the chains impairs their binding to and

246

hydrolysis by esterases. The importance of crystallinity is reflected in findings of decreasing

247

enzymatic hydrolysis rates of polyesters as their melting temperatures (Tm; temperature at

248

which

249

increase.59,100,101,104,108,112,117-119 The propensity of polyesters to form crystalline phases

250

primarily depends on their chemical structure and molecular weight distribution100,112,116,119,120

251

but is modulated by their thermal and mechanical exposure history.102,121-123

crystalline

transition

into

amorphous

phases)

and

enthalpies

of

fusion

252

A number of fungal and bacterial esterases that hydrolyze aliphatic and aliphatic-

253

aromatic co-polyesters have been identified.59,99,120,124 These esterases include cutinases and

254

lipases. The natural substrate of cutinases is cutin, a complex plant bio-polyester composed of

255

hydroxylated fatty acids125 with some chemical resemblance to synthetic polyesters. Some of

256

the identified cutinases hydrolyze semi-crystalline polyesters with high Tm, including

257

PBAT59,120 and non-biodegradable polyethylene terephthalate.126-132 Esterase activity on

258

polyesters needs to be assessed case-by-case given that several enzyme-specific factors play a

259

role (e.g., the tertiary structures and polarities of substrate binding sites and active sites that

260

can be determined from esterase crystallographic structures).

261

Knowledge gaps and research needs. While past work identified polyester properties and

262

esterase characteristics affecting enzymatic polyester hydrolysis, there are several knowledge

263

gaps related to this reaction in the context of biodegradable mulch films in soils.

264

Only a few studies have attempted to link polyester biodegradation in soils to soil esterase

265

activities.133,134 While esterase activities and PBSA-film degradation rates in eleven soils

266

correlated with the relative abundance of PBSA degrading fungi isolated from these soils, there


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was no strong direct correlation between PBSA degradation rates and soil esterase activities.134

268

This finding suggests that the substrate used in the soil esterase activity assay, p-nitrophenyl

269

valerate, was hydrolyzed by a broad set of soil esterases and, therefore, not sufficiently specific

270

to the much smaller subset of soil esterases capable of hydrolyzing PBSA. Soil esterase activity

271

assays are thus needed that employ substrates that are highly selective for soil esterases that

272

hydrolyze a specific polyester of interest. Such assays have potential to directly relate polyester

273

biodegradation rates in soils to soil esterase activities and to use soil polyester-specific esterase

274

activities to predict biodegradability of polyester-based mulch films.

275

Levels of esterase expression and secretion by polyester-colonizing microorganisms

276

remain unknown but are likely controlled by a number of factors. First, nitrogen-limited

277

microorganisms may not produce esterases given that their secretion and subsequent hydrolysis

278

of polyesters does not increase N-availability. Second, the general availability of carbon

279

substrates may regulate esterase production in colonizing microorganisms. For instance, the

280

production of a cutinase in Fusarium solani in culture studies was shown to be suppressed by

281

glucose135 (but induced by hydroxylated C16 fatty acids, the products of cutin hydrolysis.135,136)

282

Similarly, glucose and fructose suppressed the hydrolytic activity of an isolated PBAT-

283

degrading fungus grown on PBAT.76 Determining levels of and controls on esterase expression

284

and secretion requires novel analytical approaches to quantify esterase activities and to

285

characterize the extracellular proteome, including esterases, on the surface of polyester films

286

after soil incubation.

287

Past work studied enzymatic hydrolysis primarily of pure polyesters. Going forward, the

288

enzymatic hydrolyzability of commercial biodegradable mulch films — both in their pristine

289

states prior to field application as well as in weathered states after field application — needs to

290

be systematically studied for a number of reasons. First, biodegradable mulch films typically

291

contain more than one polymer. Enzymatic depolymerization of these blends may require more


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than one hydrolase and likely exhibit more complex dynamics as compared to those of single

293

polyesters.65,137 Second, most research on enzymatic polyester hydrolysis has been conducted

294

on unprocessed polyesters and not on polyester mulch films produced by blown film extrusion.

295

Because this processing step is expected to largely affect the physical properties of the

296

comprised polyesters, it is preferable to assess enzymatic hydrolyzability (and biodegradability

297

in soils) on the final, processed films.121 Third, future work needs to assess the extent to which

298

weathering of biodegradable mulch films during field application affects their enzymatic

299

hydrolyzability. In particular, the effect of photoirradiation-induced changes in the chemical

300

structures of aliphatic-aromatic co-polyesters67,138,139 on enzymatic hydrolyzability warrants

301

systematic investigation.

302

Step 3: Microbial utilization of polymer carbon. The last step in mulch film

303

biodegradation is the microbial assimilation and utilization of monomers and oligomers

304

released from the polymers through enzymatic hydrolysis. Microorganisms utilize the

305

hydrolysis products as substrates for both respiration and synthesis of biomolecules (Figure

306

2). The most direct approach to study utilization therefore is to follow the conversion of

307

polymer-derived carbon into CO2 (also referred to as ‘mineralization’) and into microbial

308

biomass.48 Indirect approaches —including monitoring of polymer mass loss, detecting shifts

309

in the polymer molecular weight distribution towards smaller masses, and visually assessing

310

polymer physical disintegration— provide no direct evidence for microbial utilization of

311

polymers during soil incubation.

312

Conceptual understanding. The mineralization of polymer carbon into CO2 during soil 140-142

313

incubations can be directly quantified by respirometric analyses of the formed CO2.

314

Respirometric data is typically plotted as the cumulative amount of formed CO2 —corrected

315

for CO2 formation in polymer free control incubations and expressed in percent of the polymer

316

carbon added to the soil— versus incubation time (depicted for three cases of mineralization


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in Figure 4). Resulting mineralization curves are the foundation of biodegradation standards

318

that stipulate conversion of a specified amount of the polymer carbon to CO2 over a defined

319

incubation time.40,141

320

Mineralization curves provide information both on the rates and extents of polymer

321

conversion to CO2. At the onset of the incubations, the mineralization curves commonly show

322

a lag phase, presumably reflecting time required for microbial colonization of the polymers

323

and the subsequent onset of enzymatic depolymerization. Incubations showing fast rates and

324

high final extents of mineralization imply extensive biodegradation of the polymers in the

325

studied soils (Figure 4, case 1). Incubations showing slow rates and incomplete final extents

326

of mineralization (Figure 4, case 2) are more difficult to interpret as slow utilization may be

327

the result of polymer properties (e.g., crystallinity), soil characteristics (e.g., nutrient

328

availability), or the incubation conditions (temperature and soil water content). Identifying

329

causes of slow utilization would require additional incubation experiments specifically

330

targeting individual system parameters hypothesized to constrain mineralization. However,

331

systematic studies on the effects of polymer- and soil-specific factors on microbial utilization

332

remain sparse.52,143 One of these studies demonstrated that polymer utilization in 19 field soils

333

increased with increasing soil nitrogen contents and with increases in the cumulative time

334

period over which temperatures at the field sites were above 10°C.52

335

Additional difficulties in the interpretation of mineralization curves arise because the

336

non-mineralized fraction remaining in soils typically is not characterized. This fraction may be

337

present both as residual polymer or associated with soil microbial biomass (Equation 1). The

338

extent to which microorganisms incorporate carbon into their biomass is described by the

339

carbon use efficiency (CUE; i.e., the ratio of carbon used for growth to the sum of carbon used

340

for growth and respiration). The CUE is species- and substrate-specific144,145 and strongly

341

dependent on incubation conditions, including nutrient availability. As a consequence, the


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342

conversion of polymer carbon into microbial biomass cannot be accurately quantified by

343

determining the increase in the amount of microbial biomass extractable from soils over the

344

course of polymer incubations.

345

346 347 348 349 350 351 352 353

Figure 4. a. Schematic of polymer mineralization experiments in soils in which the formation of carbon dioxide (CO2) is followed over time. b. Schematic of mineralization curves for three polymer-soil combinations resulting in fast and complete mineralization (case 1), slower and incomplete mineralization (case 2), and no mineralization (case 3). Based on only respirometric data, the relative contributions of remnant polymer and microbial biomass to the nonmineralized fraction in case 2 remain unknown.

354

Knowledge gaps and research needs. To characterize the nonmineralized fraction of

355

polymers in soils, it is critical to develop analytical methods that allow quantifying the amount

356

of remnant polymers in soils. Existing gravimetric analysis methods are poorly-suited because

357

they rely on completely recovering the remnant polymer from the soil and effective cleaning

358

of the polymer prior to weighing. Instead, solvent-extraction methods, such as Soxhlet and

359

accelerated solvent extraction, to recover residual polymer from soils are more promising.146,147

360

The amount of extracted polyesters could subsequently be quantified by gel permeation

361

chromatography or nuclear magnetic resonance spectroscopy. Efficient extraction methods of

362

polyesters from soils would aid the interpretation of laboratory incubation studies but also


16


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363

allow to screen for remnant polyester in field soils to which biodegradable mulch films were

364

previously applied.

365

Currently-employed incubation systems with respirometric measurements have two

366

major limitations that impair detailed mechanistic studies on polymer utilization. First, these

367

systems only allow for measure of polymer mineralization but not for the closure of mass

368

balances on the polymer carbon added to the soils.148 Second, they offer no information on the

369

incorporation of polymer-derived carbon into soil microbial biomass. These limitations can be

370

overcome by using carbon isotope-labeled instead of unlabeled polymers in soil incubations.

371

While early studies have successfully employed

372

mineralization in soils,149,150 the use of this radioisotope is often impractical due to regulations

373

and safety measures. Instead, the use of stable

374

conversion of 13C-labeled polymers into 13CO2 during soil incubations can be quantified either

375

by gas chromatography coupled to isotope ratio mass spectrometry (IRMS) or 13CO2-sensitive

376

cavity ring down spectroscopy.60,151,152 The use of

377

possibility to quantify the amount of polymer-derived 13C that remains in soils at the end of

378

incubations, for instance by soil combustion coupled to quantification of formed

379

IRMS. By combining the quantified amounts of non-mineralized 13C remaining in soils and of

380

13C

381

polymer-derived 13CO2 can be delineated from CO2 formed in the mineralization of soil organic

382

matter, mineralization studies with

383

arise if polymer addition increases or decreases the mineralization of soil organic matter (i.e.,

384

so called positive or negative ‘priming effects’).153,154

13C-

14C-labeled

polymers to follow their

labeled polymers is more viable. The

13C-labeled

polymers also opens the

mineralized, it is possible to close the mass balance on added polymer

13C-labeled

13CO

13C.

2

by

Because

polymers are not susceptible to artifacts that

385

In addition to enabling mass balance determinations, using 13C-labeled polymers further

386

enables tracking of polymer carbon into soil microbial biomass. For instance, microbial

387

biomarkers (such as phospholipid fatty acids) from soils can be extracted and their 13C-content


17

Page 19 of 31


388

can be quantified.155,156 Quantifying

389

provide valuable insights into the relative importance of bacteria and fungi in utilizing

390

polymer-derived carbon.157 Alternatively, the incorporation of polymer-derived

391

polymer-colonizing microorganisms can be directly imaged using

392

microscopy techniques, including Nanoscale secondary ion mass spectrometry (NanoSIMS) or

393

Raman microspectroscopy.158-160 A recent study used NanoSIMS to unequivocally demonstrate

394

incorporation of

395

microorganisms.60

396

13C

13C

in signature bacterial and fungal biomarkers would

13C-isotope

13C

into

selective

carbon from labeled PBAT into the biomass of colonizing soil

The analytical advances outlined above —involving soil incubation of both unlabeled 13C-labeled

397

and

polymers— would enable systematic studies on the effects of polymer

398

properties and soil characteristics on the CUE of microbial polymer degraders. Determining

399

the effects of N availability on CUEs and on mineralization rates and extents is particularly

400

pertinent. CUEs of soil microorganisms are known to decrease from around 0.3-0.5 for

401

substrates with low C:N to much smaller values for substrates with high C:N ratios.161 Because

402

polymers used in biodegradable mulch films (Figure 1) fall into the latter category, increasing

403

N availability in soils likely increases CUEs and overall polymer biodegradation.

404

Concluding thoughts

405

Using biodegradable mulch films instead of conventional PE-based films promised to

406

improve the sustainably of agricultural food production by overcoming deleterious economic

407

and ecological impacts resulting from the accumulation of remnant PE films in agricultural

408

soils. To ensure the safe application of biodegradable mulch films —including their desired

409

biodegradation in soils— requests that their fate in soils is well studied and understood at a

410

mechanistic level. This perspective highlights research and analytical developments required

411

to advance a more fundamental understanding of the key steps involved in mulch film

412

biodegradation in soils. This understanding will be beneficial to establishing transferability of


18


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413

results from polymer biodegradation studied in laboratory batch and mesocosm incubations to

414

biodegradation in agricultural soils directly in the field, determining the relative importance of

415

(specific) soil fungi and bacteria in film biodegradation, assessing and adjusting mulch film

416

biodegradation in field soils, creating tools to predict biodegradability based on site-dependent

417

soil characteristics and climatic factors, and to further refining standards and test methods for

418

the biodegradation of mulch films in soils.

419

Acknowledgements

420

M.S. thanks Gordon Getzinger, Michael Zumstein, and Taylor Nelson for critical

421

feedback on the manuscript and Rebekka Baumgartner, Michael Zumstein, Taylor Nelson,

422

Mattia Cerri, Kristopher McNeill (all ETH) and Hans-Peter Kohler (Eawag) for an inspiring

423

collaboration over the past years on the fate of biodegradable polymers in soils. Parts of the

424

content of this perspective reflects discussions with them. The SEM images in Figure 3 were

425

collected by Michael Zumstein using equipment maintained by the Center for Microscopy and

426

Image Analysis, University of Zurich.

427 428 429 430 431 432 433 434 435 436 437 438 439 440 441 442 443 444 445 446 447

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Short Bio Michael Sander

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Michael Sander is a senior scientist in Environmental Chemistry at the Swiss Federal Institute of Technology in Zurich (ETHZ), Switzerland. Michael Sander received his M.S. degree in Environmental Sciences from the University of Bayreuth, Germany, in 2000 and his Ph.D. degree in Chemical Engineering (Environmental Engineering Program) from Yale University, USA, in 2005. Following postdoctoral research from 2005 to 2007 in Environmental Chemistry at ETHZ, Michael Sander was promoted to research group leader in 2008 and to his current position in 2017. Michael Sander’s research group is active in three areas: environmental organic chemistry of micropollutants, redox (biogeo)chemistry, and environmental macromolecular chemistry. Research in the latter area is directed towards advancing a fundamental understanding of the environmental fate and activity of (bio)macromolecules, including enzymes, plant-incorporated protectants, viruses, and biodegradable synthetic organic polymers. The work from Michael Sander’s research group has received several awards including two Best Paper Awards from Environmental Science & Technology (categories Environmental Technology in 2016 and Environmental Science in 2009) and an Excellence in Review Award from the same journal in 2013.


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Picture Michael Sander


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Biodegradation of Polymeric Mulch Films in Agricultural Soils

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