Abundant Nonprotonated Aromatic and Oxygen-bonded Carbons

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Abundant Nonprotonated Aromatic and Oxygen-bonded Carbons Make Humic Substances Distinct from Biopolymers Xiaoyan Cao, and Klaus Schmidt-Rohr Environ. Sci. Technol. Lett., Just Accepted Manuscript • DOI: 10.1021/acs.estlett.8b00107 • Publication Date (Web): 03 Apr 2018 Downloaded from http://pubs.acs.org on April 5, 2018

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Environmental Science & Technology Letters

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Abundant Nonprotonated Aromatic and Oxygen-bonded Carbons Make Humic

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Substances Distinct from Biopolymers

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Xiaoyan Cao, Klaus Schmidt-Rohr*

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Department of Chemistry, Brandeis University, Waltham, Massachusetts 02453, United States

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* Corresponding author: E-mail: [email protected]. Phone: 781-736-2520. Fax: 781-736-2516.

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1 ACS Paragon Plus Environment


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Abstract

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The nature of humic substances (HSs) extracted from soil organic matter and their distinction

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from degrading plant material has recently been called into question. Quantitative solid-state

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show that HSs are distinct from common biopolymers, refuting claims to the contrary made

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based on 1H-detected NMR spectra of the same standard peat humic acid as studied here.

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Aromatic carbons not bonded to hydrogen or oxygen, which are rare (≤ 10%) in common

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biomolecules, constitute a large fraction (28%-33%) of prairie soil and peat humic acids.

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Oxygen-bonded nonprotonated carbons such as aryl ketones, nonprotonated alkyl C-O, and

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abundant COOH groups are also characteristic of humic and fulvic acids but not observed in

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plant biopolymers. These distinctive structural differences between HSs and biopolymers

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challenge the recent proposal that HSs are unremarkable intermediates in a continuum of

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degrading biomolecules.

C nuclear magnetic resonance (NMR) analyses, in good agreement with elemental analysis,

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Introduction

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Globally, soil organic matter (SOM) holds more carbon than the atmosphere and terrestrial

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vegetation combined. It is a heterogeneous mixture of organic compounds mostly of plant and

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microbial origins in various stages of transformation. Significantly transformed organic matter

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that does not fall into any of the discrete classes of biomolecules (carbohydrates, lignin,

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proteins, amino-sugars, etc.) has been designated as humic substances (HSs).1-3 Operationally,

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HSs are usually extracted at high pH and classified into three categories based on their

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solubility: acid-soluble fulvic acid (FA), acid-insoluble humic acid (HA), and base-insoluble

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humin. Despite over a century of efforts to structurally define HSs, views on the nature of

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SOM and HSs have remained widely divergent.2-5

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A recent high-profile study4 has questioned the validity of the very concept of “humic

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substances”, arguing that it is poorly defined and that alkaline soil extracts consist simply of

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degrading biomolecules. In support of this view, a solution 1H nuclear magnetic resonance

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(NMR) study5 of a peat humic acid was quoted as “direct proof that the chemical makeup of

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‘humic substances’ can be explained as resulting from mixtures of known plant and microbial

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compounds”.4 Indeed, Ref. 5 claimed that HSs are predominantly mixtures of plant and

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microbially derived components in four chemical categories (protein, lignin, carbohydrate, and

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aliphatic biopolymers). However, the 1H NMR-based analyses used in Ref. 5 could only detect

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carbons bonded to hydrogen and focused on abundant small molecules because these sites

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generate the sharpest NMR lines and strongest surviving signals in the two-dimensional (2D)

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solution NMR spectra shown. Due to these limitations, the conclusions drawn about the

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chemical makeup of base-extracted HSs in Ref. 5 remain open to debate.

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Here we employ quantitative solid-state

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C NMR methods, which can detect all



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carbons regardless of solubility and molar mass, to comprehensively investigate the structure

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of standard HS fractions from the International Humic Substances Society (IHSS), including

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the Florida peat HA studied in Ref. 5. We also present quantitative spectra of nonprotonated

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carbons (i.e. carbons not bonded to hydrogen), which represent a major structural component

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of many humic acids but were unfortunately missing from the 1H-detected NMR spectra of Ref.

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5. By comparing the humic-acid spectra with spectra of common plant biopolymers, we test

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the hypothesis that abundant carbon-bonded aromatics and unique oxygen-bonded

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nonprotonated carbons, which are not significantly detected in the previous 1H-NMR based

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study, make extracted HSs distinct from identifiable biomolecules.

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Materials and Methods

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Three standard samples from the International Humic Substances Society (IHSS) were

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studied. The Florida peat HA standard (#1S103H) and Elliott soil HA standard (#1S102H)

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were kindly provided by Dr. Paul R. Bloom, University of Minnesota, St. Paul. The Elliott soil

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(fine, illitic, mesic Aquic Argiudoll; USDA Taxonomy) is typical of the fertile prairie soils of

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Indiana, Illinois, and Iowa.6 The Florida peat FA standard (#2S103F) was purchased from the

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IHSS. Microcrystalline cellulose (Sigma-Aldrich) and commercial cork were used as

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purchased. The 13C-enriched switchgrass and beech woody stems were purchased from Isolife

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(Wageningen, The Netherlands). Milled wood lignin from loblolly pine was provided by Dr.

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Kevin M. Holtman,7 13C-enriched GB1 protein by Dr. Chad Rienstra,8 cultured Synechococcus

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cell material by Dr. Margaret R. Mulholland,9 and maple wood char (from pyrolysis at 400 °C)

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by Dr. Joseph J. Pignatello.10


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All NMR experiments were performed at 100 MHz for 13C and 400 MHz for 1H using a

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Bruker Avance 400 spectrometer equipped with a 4-mm double-resonance probe head. The 13C

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chemical shifts were referenced to tetramethylsilane, using the COO resonance of 13C1-labeled

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glycine in the α-modification at 176.46 ppm as a secondary reference. Quantitative 13C NMR

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spectra were measured using direct polarization (DP), 14-kHz magic-angle spinning (MAS),

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and high-power decoupling,11, 12 with recycle delays of 15 s for Florida peat HA, 35 s for

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Elliott soil HA, 66 s for Florida peat FA, 100 s for switchgrass and beech woody stems, and 10

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s for Synechococcus cells. Nearly quantitative spectra with superior signal-to-noise ratio were

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obtained with composite-pulse multiple cross polarization (multiCP)13 with a repolarization

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delay of 1 s and 10 rampCP blocks of 1.1 ms duration, plus a final 0.55 ms CP period. A Hahn

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echo (at 2τ = 0.14 ms) was generated before detection to avoid baseline distortions arising from

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pulse dead time.12 Corresponding spectra of nonprotonated carbons and mobile segments were

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obtained similarly after recoupled 13C-1H dipolar dephasing of 68-µs duration.12

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Results and Discussion

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Figure 1a shows a nearly quantitative multiCP

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same standard HA as studied in Ref. 5. The spectrum is dominated by peaks of aromatic

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carbons, which were observed only at low intensity in the 1H-detected spectra of Ref. 5. The

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corresponding quantitative but noisier direct-polarization spectrum is shown in Figure S1a. The

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solid-state NMR spectrum (Figure 1a) closely resembles a solution 13C NMR spectrum of the

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same standard material.14 By contrast, the 1H NMR spectrum of Ref. 5 does not closely match

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previously published 1H spectra,14 which may be due to the solubilizing pretreatment applied in

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Ref. 5. Furthermore, the high abundance of peaks due to nonprotonated aromatic and carboxyl

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C NMR spectrum of the IHSS peat HA, the



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carbons is a typical feature observed in 13C NMR spectra of soil humic acids of various origins

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and locations.15-19 Figure 1b shows the 13C NMR spectra of four common biopolymers, similar

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to those used in Ref. 5. The positions and relative intensities of the peak in the biopolymer

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spectra are in good agreement with the literature.8, 20-22 Superposition of the biopolymer spectra

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in an attempt to match the HA spectrum unavoidably leaves major discrepancies, see Figure

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S1b, in particular in the aromatic carbon region.

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The nonprotonated carbons in the humic acid can be observed quantitatively and

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selectively at > 60 ppm in the spectrum of Figure 2a after recoupled dipolar dephasing,12 which

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is a useful if underutilized spectral-editing technique for soil organic matter.12, 23, 24 Note that

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all of the nonprotonated carbon sites, which according to their signal area account for

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approximately 60% of all C (Table S1), were invisible in the 1H-detected spectra shown in Ref.

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5. The spectrum shows pronounced discrepancies with the corresponding spectra of the

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biopolymers (Figure 2b), which contain little signal intensity at and to the right of the humic

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acid aromatic signal maximum near 130 ppm. This refutes the claim that “nearly all of the

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NMR signals in traditional HS fractions could be assigned to intact and degrading

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biopolymers.”5 The agreement between the CHn-selective 1H-detected NMR spectra of the peat

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humic acid and the spectra of biopolymers5 shows that biopolymer residues are present in

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SOM at some level, but this is not a surprising proposition; it is widely accepted by soil

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scientists2, 25 that the base-extracted SOM is HS-rich, but also contains some relatively fresh

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plant or microbial biomolecules.2

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The four biopolymer types considered here and in Ref. 5 have H:C atomic ratios

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ranging between 0.85 and 2.26 This range is not consistent with the H:C ratio of 0.8 in the IHSS

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peat HA,27 confirming that this humic acid is distinct from plant biopolymers. On the other


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hand, the atomic ratio of H:C = 0.82 ± 0.07 calculated from the peak integrals15 of the

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quantitative, aromatic-rich

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reported H:C ratio, as does the (O+N)/C ratio (0.53 ± 0.06 from NMR vs. 0.55 measured27).

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These data, which are consistent with a previous NMR study of several humic acids,15 validate

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the reliability of our NMR analysis.

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C NMR spectra in Figures 1a and 2a agrees quite well with the

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Looking beyond the nearly pure biopolymers shown in Figures 1 and 2, we can also

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compare the spectra of HSs and plant or microbial biomass (Figures 3 and S2). Figure 3 shows

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quantitative

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microbial biomass, with a significant protein contribution. The spectra of their nonprotonated

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carbons, shown as thick lines, show little signal at < 130 ppm and > 185 ppm, unlike the humic

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acid in Figure 2. The much higher aromatic fraction in the humic acid, even compared to wood,

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is striking and highlights the recalcitrance of aromatic structures in SOM. Further, the

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pronounced mismatch between the spectra of microbial biomass and HSs casts doubt on the

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hypothesis that microbial residues are the main precursors of SOM.28

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C NMR spectra of grass and wood, dominated by carbohydrate signals, and of

The large fraction of nonprotonated aromatics observed in the

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C spectrum of IHSS 13

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peat HA is significant in other HSs as well. For example, the nearly quantitative

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spectra of IHSS peat FA and Elliot soil HA (Figure 3) are dominated by signals of

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nonprotonated aromatics and carboxyl groups that are clearly identified after dipolar dephasing

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(thick lines in Figure 3). The peak maximum of the nonprotonated carbons near 130 ppm

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cannot arise from aromatic carbons within one or two bonds from oxygen, such as those found

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in polyphenols, since these usually resonate at > 138 ppm or < 125 ppm.29 Instead, it is typical

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of condensed or COO-substituted aromatic rings. Long-range 1H-13C dipolar dephasing has

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indeed shown that many of the nonprotonated aromatic carbons in peat and Mollisol HAs are


C NMR


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further from hydrogen than in lignin,18,

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formation from biopolymer precursors requires new bond formation. The aromatic carbons not

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bonded to hydrogen or oxygen constitute large fractions (28% and 33%, Table S1) of peat and

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prairie soil humic acids, respectively, while they are rare (≤ 10%) in common biopolymers.

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The nonprotonated aromatics resonating at 130 ppm may be indicative of oxidized char

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residues, which have been shown to be a major component (up to 50%) of Mollisols and of

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their humic acids (~70%),18 probably having been generated by recurring prairie fires.

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Evidence of pyrogenic or black carbon has also been found in other soil types and sediments,17

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in particular volcanic ash soils,31-34 again likely arising from vegetation fires.35 For reference,

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Figure 4c shows the spectrum of maple wood char made by pyrolysis at 400 oC, which shows

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nonprotonated aromatics in a similar proportion as the HSs but fewer signals associated with

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oxygen.

indicating condensation of aromatic rings; their

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The major nonprotonated aromatic fraction observed here challenges the assumption in

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Ref. 5 that black carbon is a less abundant biochemical group in HSs than are four categories

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of biopolymers (protein, lignin, carbohydrate, and lipids/waxes). It is also interesting to note

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that the predominance of condensed aromatic structures in extracted HSs provides a sufficient

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explanation for the “dark colour” of alkaline soil extracts listed at the top of Figure 3 of Ref. 4;

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the proposed explanations in terms of conjugated bonds, which would be prone to oxidation, or

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“degradation of pigments”,4 do not account for the large fraction of condensed aromatics

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observed here.

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In the spectrum of the peat HA (Figures 1 and S1), signals of aromatic carbons near

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oxygen at 150 and around 120 ppm are also quite pronounced and could be attributed to

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reactions primarily of lignin in soil during humification.3,

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It is noteworthy that lignin

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contains hardly any nonprotonated carbons resonating below 125 ppm, see Figure 2, while

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such aromatic carbons are abundant in humic acids. These sites must be bonded to three other

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carbons, according to spectral editing, and be separated from one oxygen atom by exactly two

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bonds, according to their chemical shift,29 which indicates that new carbon-carbon bond

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formation has occurred during the transformation of the biopolymer precursors, in particular

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lignin, to more complex molecules in the humic acid, which is consistent with the concept of

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humification.2

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The spectra of peat humic and fulvic acids also show a distinct peak near 200 ppm,

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which is assigned to ketones bonded to an aromatic and an alkyl carbon,38-40 which are not

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common in biopolymers (Figures 1-4). Aromatic ketones have also been detected in aquatic

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HSs and have been proposed to arise from photo-Fries rearrangement of phenolic esters or

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photo oxidation of aryl-aliphatic hydrocarbons.39 Additionally, the signal intensity near 185

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ppm may be assigned to quinones, which have been identified in soil HSs by 15N NMR after

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derivatization.40 The NMR signal near 85 ppm after dipolar dephasing, seen most distinctly for

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the peat fulvic acid, is also unique to spectra of HSs (Figures 2 and 4). Based on its chemical

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shift, this type of nonprotonated carbon must be sp3-hybridized and bonded to oxygen, i.e.

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bonded to three carbon atoms and one oxygen (“quaternary” alkyl C-O, “OCq”). Such an OCq

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structure is absent from common plant biopolymers but a significant component of dissolved

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organic matter in aquatic environments.41-43 Given that the OCq site has three carbon neighbors,

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new carbon–carbon bond formation is again likely. These C=O and OCq structures, which are

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easily recognized due to their distinctive chemical shifts, can be considered as the “tip of the

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iceberg” of pronounced structural transformations of biomolecules in soil that are consistent

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with the concept of humification.3 Finally, carboxylic acid groups showing a strong peak near



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171 ppm are greatly enriched in humic and fulvic acids relative to biopolymers, and are the

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namesakes of humic and fulvic acids.

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In summary, the present study has demonstrated that extracted HSs are clearly distinct

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from plant and microbial biopolymers in that they contain abundant condensed aromatics,

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nonprotonated aromatic carbons near oxygen, as well as oxygen-bonded nonprotonated

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carbons such as ketone, OCq, and abundant carboxylic acid groups. This refutes a previous

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conclusion that the NMR signals of HSs could be assigned nearly exclusively to intact and

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degrading biopolymers,5 and challenges the recent proposal that alkaline extracts are

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unremarkable intermediates in a continuum of degrading plant residues.4

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Figure 1. (a) Nearly quantitative multiCP

C NMR spectrum of IHSS peat humic acid,

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compared with (b) spectra of lignin, protein, cellulose, and cork (dominated by polymethylene-

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rich suberin resonating near 32 ppm), all scaled to the same maximum peak height.

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Figure 2. (a) Dipolar dephased multiCP 13C NMR spectrum of nonprotonated carbons (at > 60

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ppm) and mobile CHn groups in IHSS peat HA. Note that most of these signals were absent

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from 1H-detected solution NMR spectra in Ref. 5, even from the 2D heteronuclear single

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quantum coherence spectrum with its 13C dimension. (b) Dipolar dephased 13C NMR spectra of

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the same four biopolymers as in Figure 1b. In the spectral ranges marked by shading, the

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biopolymer spectra do not provide signal intensity to match the HA spectrum.


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C NMR spectra of uniformly

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Figure 3. Quantitative

C-enriched biomass: (a) switchgrass,

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(b) beech wood, and (c) cyanobacteria (Synechococcus). Thick lines: spectra of nonprotonated

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and mobile C. Within the dashed lines as in Figure 2, little nonprotonated carbon signal is

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detected. More spectra of biomass are shown in Figure S2.

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Figure 4. Nearly quantitative multiCP 13C NMR spectra of (a) IHSS peat fulvic acid, (b) IHSS

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Elliot soil humic acid, and (c) maple wood char made by pyrolysis at 400 oC. Thick lines:

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spectrum after dipolar dephasing. Corresponding direct-polarization spectra are shown in

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Figure S3.

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Notes

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The authors declare no competing financial interest.

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Supporting Information

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Quantitative direct-polarization 13C NMR spectra of extracted HSs, a sum spectrum of the four

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biopolymers, description and spectra of additional biomass and microbial samples, and a table

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summarizing the functional group composition of HSs, char, and biomass samples. This

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material is available free of charge via the Internet at http://pubs.acs.org.

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Acknowledgments. The authors thank the IHSS for making standard humic substances

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available, which allows for reliable comparison of results from different laboratories.

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Stimulating discussions with Drs. Jingdong Mao and Daniel C. Olk are gratefully

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acknowledged.

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References

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26. Visser, S. A., Application of Van Krevelen's graphical-statistical method for the study of aquatic humic material. Environ. Sci. Technol. 1983, 17, 412-417. 27. International Humic Substances Society. Elemental Compositions and Stable Isotopic Ratios of IHSS Samples. http://humic-substances.org/elemental-compositions-and-stableisotopic-ratios-of-ihss-samples/ (date accessed February 20, 2018). 28. Cotrufo, M. F.; Wallenstein, M. D.; Boot, C. M.; Denef, K.; Paul, E., The Microbial Efficiency‐Matrix Stabilization (MEMS) framework integrates plant litter decomposition with soil organic matter stabilization: do labile plant inputs form stable soil organic matter? Global Change Biol. 2013, 19, 988-995. 29. Badertscher, M.; Bühlmann, P.; Pretsch, E., Structure determination of organic compounds. Springer Berlin Heidelberg: 2009. 30. Mao, J.-D.; Schmidt-Rohr, K., Recoupled long-range C–H dipolar dephasing in solidstate NMR, and its use for spectral selection of fused aromatic rings. J. Magn. Reson. 2003, 162, 217-227. 31. Shindo, H.; Matsui, Y.; Higashi, T., A possible source of humic acids in volcanic ash soils in Japan-charred residue of Miscanthus sinensis. Soil Sci. 1986, 141, 84-87. 32. Kramer, R. W.; Kujawinski, E. B.; Hatcher, P. G., Identification of black carbon derived structures in a volcanic ash soil humic acid by Fourier transform ion cyclotron resonance mass spectrometry. Environ. Sci. Technol. 2004, 38, 3387-3395. 33. Ikeya, K.; Sleighter, R. L.; Hatcher, P. G.; Watanabe, A., Characterization of the chemical composition of soil humic acids using Fourier transform ion cyclotron resonance mass spectrometry. Geochim. Cosmochim. Acta 2015, 153, 169-182. 34. Ikeya, K.; Hikage, T.; Arai, S.; Watanabe, A., Size distribution of condensed aromatic rings in various soil humic acids. Org. Geochem. 2011, 42, 55-61. 35. Schmidt, M. W.; Noack, A. G., Black carbon in soils and sediments: analysis, distribution, implications, and current challenges. Global Biogeochem. Cycles 2000, 14, 777793. 36. DiDonato, N.; Chen, H.; Waggoner, D.; Hatcher, P. G., Potential origin and formation for molecular components of humic acids in soils. Geochim. Cosmochim. Acta 2016, 178, 210222. 37. Leenheer, J. A., Systematic approaches to comprehensive analyses of natural organic matter. Ann. Environ. Sci. 2009, 3, 1-130. 38. Johnson, R. L.; Anderson, J. M.; Shanks, B. H.; Fang, X.; Hong, M.; Schmidt-Rohr, K., Spectrally edited 2D 13C-13C NMR spectra without diagonal ridge for characterizing 13Cenriched low-temperature carbon materials. J. Magn. Reson. 2013, 234, 112-124. 39. Leenheer, J.; Wilson, M.; Malcolm, R., Presence and potential significance of aromaticketone groups in aquatic humic substances. Org. Geochem. 1987, 11, 273-280. 40. Thorn, K. A.; Arterburn, J. B.; Mikita, M. A., 15N and 13C NMR investigation of hydroxylamine-derivatized humic substances. Environ. Sci. Technol. 1992, 26, 107-116. 41. Cao, X.; Aiken, G. R.; Butler, K.; Mao, J.; Schmidt-Rohr, K., Comparison of the chemical composition of dissolved organic matter in three lakes in Minnesota, USA. Environ. Sci. Technol. 2018, 52, 1747-1755. 42. Cao, X.; Aiken, G. R.; Butler, K. D.; Huntington, T. G.; Balch, W. M.; Mao, J.; Schmidt-Rohr, K., Evidence for major input of riverine organic matter into the ocean. Org. Geochem. 2018, 116, 62-76.



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43. Cao, X.; Aiken, G. R.; Spencer, R. G.; Butler, K.; Mao, J.; Schmidt-Rohr, K., Novel insights from NMR spectroscopy into seasonal changes in the composition of dissolved organic matter exported to the Bering Sea by the Yukon River. Geochim. Cosmochim. Acta 2016, 181, 72-88.

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