University of California Santa Cruz

UNIVERSITY OF CALIFORNIA SANTA CRUZ

PHOSPHORUS CYCLING IN THE DEEP SEDIMENTARY SUBSEAFLOOR ENVIRONMENT

A dissertation submitted in partial satisfaction of the requirement for the degree of

DOCTOR OF PHILOSOPHY

EARTH SCIENCES

Delphine Defforey

December 2016

This dissertation of Delphine Defforey is approved:

______Adina Paytan, Research Scientist

______Professor Slawek Tulaczyk, Chair

______Professor Andrew T. Fisher

______Assistant Professor Jason B. Sylvan

______Tyrus Miller Vice Provost and Dean of Graduate Studies

TABLE OF CONTENTS

Lits of figures and tables v

Abstract vi

Acknowledgements and dedications viii

Introduction 1

Chapter 1. A new solution 31P NMR sample preparation scheme for 6 marine sediments

Abstract 7 1.1 Introduction 8 1.2 Materials and procedures 1.2.1 Materials 12 1.2.2 Extraction procedure 13 1.3 Assessment 1.3.1 Evaluation of extraction procedure on natural samples 19 1.3.2 Pretreatment effects on sample chemistry 21 1.4 Discussion 23 1.5 Comments and recommendations 28 1.6 References 30

Chapter 2. Phosphorus geochemistry of sediments in ridge-flank systems 48

Abstract 49 1 Introduction 1.1 Motivation and project goal 50 1.2 Field site locations 1.2.1 Mid-Atlantic Ridge Flank 52 1.2.2 Juan de Fuca Ridge Flank 53 2 Materials and Methods 2.1 Sample collection and storage 54 2.2 Sediment sequential extraction 55 2.3 Supernatant P analyses 56 2.4 Nannofossil biostratigraphy 57 2.5 Phosphorus accumulation rates 57 2.6 Statistical analyses 58 3 Results

iii 3.1 North Pond sedimentary P geochemistry 59 3.2 Grizzly Bare seamount sedimentary P geochemistry 61 4 Discussion 4.1 North Pond sedimentary P cycling 63 4.2 Grizzly Bare solid-phase P cycling 66 5 Summary and conclusions 68 6 References 70

Chapter 3. Potential phosphorus uptake mechanisms in the deep 89 sedimentary biosphere

Abstract 90 1 Introduction 91 2 Materials and Methods 2.1 Sample collection and storage 93 2.2 DNA extraction 94 2.3 Bioinformatic analyses 95 2.4 16S rRNA phylogenetic tree 96 2.5 Solution 31P NMR extraction 97 2.6 Solution 31P NMR analysis and post-experimental processing 98 2.7 Supernatant P analyses 99 3 Results and Discussion 100 4 References 109

Conclusions 126

iv FIGURES AND TABLES

Chapter 1. A new solution 31P NMR sample preparation scheme for marine sediments

1–1 Stepwise extraction scheme 37 1–2 P concentrations of sediment samples used in this study 38 1–3 31P NMR spectra of Equatorial Pacific sediment samples 39 1–4 31P NMR spectra of California Margin sediment samples 40 1–5 Pretreatment supernatant phosphorus concentrations 41 1–6 31P NMR spectra of CDB degradation test 42 1–7 31P NMR spectra of sodium acetate degradation test 43 T1–1 Sample characteristics 44 T1–2 Phosphorus yields for each extraction step 45 T1–3 Solution 31P NMR extract chemistry 46 T1–4 Measurement uncertainties associated with each extraction step 47

Chapter 2. Phosphorus geochemistry of sediments in ridge-flank systems

2–1 Location of the study’s field sites 75 2–2 North Pond SEDEX solid-phase P concentrations 76 2–3 North Pond P accumulation rates through time 77 2–4 Grizzly Bare SEDEX solid-phase P concentrations 78 T2–1 Hole locations and specifications 79 T2–2 SEDEX method uncertainty assessment 80 T2–3 Site U1363 (IODP X327) sediment chemistry 81 S2–1 North Pond nannofossil biostratigraphy 82 S2–2 North Pond total sedimentary P and accumulation rates 87 S2–3 Grizzly Bare total sedimentary P and accumulation rates 88

Chapter 3. Potential phosphorus uptake mechanisms in the deep sedimentary biosphere

3–1 Location of the study’s field site at North Pond 115 3–2 North Pond solution 31P NMR spectra 116 3–3 Pretreatment supernatant P concentrations 117 T3–1 P genes present in North Pond sediments 118 T3–2 P compounds detected with solution 31P NMR 122 T3–3 Solution 31P NMR extraction yields 123 T3–4 Solution 31P NMR extract chemistry 124 S3–1 16S rRNA phylogenetic tree 125

ABSTRACT

Phosphorus cycling in the deep sedimentary subseafloor environment

Delphine Defforey

The deep subseafloor environment is dominantly a prokaryotic habitat, with active microbial communities extending into the upper oceanic crust and affecting global biogeochemical cycles. The focus of this thesis is on phosphorus (P) biogeochemical cycling in marine sediments, particularly carbon-depleted open ocean sediments, which represent approximately 70% of the seafloor. The sediment samples used in each study are from different ocean basins and span a wide range of redox conditions, sediment types, basement ages, hydrothermal settings and P sources to the seabed, and offer valuable insights into how these parameters affect sedimentary P composition, as well as biotic and abiotic processes altering solid-phase P. Each study in this thesis provides new insights into sediment P biogeochemistry by using an array of interdisciplinary tools. In the first study, a new method for preparing sediment samples for solution 31P nuclear magnetic resonance spectroscopy (31P

NMR) is presented that enhances the use of this tool with organic matter–depleted sediments by amplifying the signal of low abundance P forms, which include organic

P compounds. This is achieved with a two–step pretreatment prior to the solution 31P

NMR extraction that quantitatively removes orthophosphate, thus increasing the

vi signal of low abundance P forms relative to orthophosphate without the use of extremely long experiments. This new method allows the use of this tool on open- ocean sediments and substantially improves our understanding of the dynamics of organic P in this environment. Results from the second study shed light on sedimentary P geochemistry of ridge-flank systems, which are regions far from the magmatic influence of seafloor spreading, including areas that are coming under the influence of (but not yet undergoing) subduction. This study focuses on two ridge- flanks, the Mid-Atlantic Ridge flank and the Juan de Fuca Ridge flank, where active hydrothermal circulation in young oceanic crust affects not only microorganisms in sediments and upper basaltic crust, but also chemical fluxes in this setting (especially at the Mid-Atlantic Ridge flank). Sequential extractions of sediments from these two well-studied sites reveal that P is mainly present in mineral phases, which vary in abundance based on sediment redox conditions and P source to the seafloor. This study also highlights the persistence of organic P throughout the sediment column in these open ocean sediments. The third study uses a combined approach of metagenomic analyses and solution 31P NMR to explore potential microbial P uptake mechanisms and P substrates for the deep biosphere. Results from this study show that organic P forms in oligotrophic sediments beneath the North Atlantic Gyre are diverse, and decrease in relative abundance with depth and changing redox conditions, thus supporting the putative P uptake pathways identified with the metagenome.

vii

ACKNOWLEDGEMENTS AND DEDICATION

I would like to thank my advisor, Adina Paytan, for her unwavering support over the course of my degree and for the opportunity to work on such an interesting set of projects. I would also like to thank the members of my reading committee for their contributions and support, and acknowledge the collaborators involved in this work:

Dr. Barbara Cade-Menun, Dr. Benjamin Tully, Dr. Jason B. Sylvan, Dr. Leah LeVay and Dr. Brandi Kiel Reese. I am also very thankful for the help and support from Dr.

Corey Liu, Rob Franks, members of the Paytan lab and the many undergraduate students who helped me with lab work over the years, including: Michael Kong,

Caitriona Berger and Natalie Zimdahl.

In dedication to my grandparents: Denis and Marguerite Defforey,

Jeanine and Jacques Charlier.

viii INTRODUCTION

Phosphorus is a critical nutrient for life, as it provides the phosphate-ester backbone of DNA and RNA, and plays a crucial role in the transmission of chemical energy by the ATP molecule. P is also a structural constituent in many cell components such as phosphoproteins, and phospholipids in cell membranes. Furthermore, it is involved in many cellular processes through the mechanism of phosphorylation, which alters the function and activity of proteins. Hence, P availability plays a substantial role in regulating primary productivity in some marine systems and, over geological timescales, in the world’s oceans [Krom et al., 1991; Benitez-Nelson, 2000; Wu et al.,

2000]. This has profound implications on ocean and atmospheric chemistry.

Despite the extensive knowledge of the global P cycle, some aspects of its marine component remain understudied. Economic and scientific interests in phosphorites

(phosphate-rich deposits) have resulted in many studies focused on sedimentary P geochemical forms, sinks and rates of burial [Delaney, 1998]. Despite this, little is known about P cycling in deep subseafloor sediments, P species fueling microbial life in this environment and how the composition and concentration of P geochemical forms change with different environmental conditions. We also know little about the influence of microbes on deep sediment P diagenetic processes.

1 The deep-subseafloor environment is estimated to contain up to 1% Earth’s total biomass, and includes significant prokaryotic populations at depths greater than a kilometer [Roussel et al., 2008; Kallmeyer et al., 2012]. The existence and activity of these microbial populations may have implications on global biogeochemical cycles and our understanding of the limits of life. Microorganisms can promote rock and mineral weathering by taking up mineral constituents using metabolic byproducts such as inorganic/organic acids, as well as ligands [Ehrlich, 1998; Banfield et al.,

1999]. Microbial biofilms can also modify local redox conditions on the surface of mineral grains, and amplify the dissolution of minerals [Ehrlich, 1998]. Furthermore, microbes play a role in the authigenic/diagenetic formation of certain minerals/sedimentary rocks [Hirschler et al., 1990a, 1990b; Ehrlich, 1998]. Lastly, deep subseafloor microbes play a significant role in controlling the chemical composition of the deep-ocean and atmosphere by selectively degrading organic matter through metabolic respiration [Parkes et al., 1994]. For these reasons, a more thorough understanding of the role P plays in this system is required.

This thesis investigates sedimentary phosphorus (P) biogeochemistry in the open ocean using a multidisciplinary approach. The first chapter offers a new method for preparing marine sediment samples for solution 31P nuclear magnetic resonance spectroscopy (31P NMR) in order to determine the chemical structure of P-bearing compounds, both inorganic and organic, in open ocean sediments. This method improves on previous work by enhancing the use of solution 31P NMR as an effective

2 tool to study organic P in sediments with low organic P content (less than 10% of total P), which represent a large fraction of marine sediments. The second study explores solid-phase P geochemistry of sediments in ridge-flank environments through the use of sediment sequential extractions, which categorize sedimentary P into five operationally-defined reservoirs: (1) loosely-sorbed P; (2) ferric Fe-bound P;

(3) authigenic carbonate fluorapatite, biogenic apatite and calcium carbonate-

associated P (CaCO3-bound P); (4) detrital apatite and (5) organic P [Ruttenberg,

1992; Ruttenberg et al., 2009]. Two field sites –one on the Mid-Atlantic Ridge flank, the other on the Juan de Fuca Ridge flank– were chosen based on their location in separate ocean basins, differing hydrothermal activity, basement age, and redox conditions. This allowed for valuable insights into changes in composition and concentration of P species as a result of different environmental conditions, such as organic matter content, redox conditions, burial depth and sediment composition. The third chapter uses both metagenomic analyses and 31P NMR (using the method developed in chapter 1) to investigate potential microbial P uptake mechanisms at play in this environment, as well as P substrates available to fuel the deep sedimentary biosphere. Results from this innovative study suggest that that the deep biosphere may play a role in the gradual breakdown of inositol and sugar phosphates, as well as reduced P compounds and polyphosphates in low organic matter sediments.

Each study described herein contributes to our understanding of P biogeochemistry in open ocean sediments, and this work constitutes a significant advance in our

3 understanding of the nature of sedimentary P and the role it plays in fueling microbial life in the deep sedimentary subseafloor environment.

References

Banfield, J. F., W. W. Barker, S. A. Welch, and A. Taunton (1999), Biological impact on mineral dissolution: Application of the lichen model to understanding mineral weathering in the rhizosphere, Proc. Natl. Acad. Sci., 96(7), 3404– 3411, doi:10.1073/pnas.96.7.3404.

Benitez-Nelson, C. R. (2000), The biogeochemical cycling of phosphorus in marine systems, Earth-Sci. Rev., 51(1), 109–135, doi:10.1016/S0012-8252(00)00018- 0.

Delaney, M. L. (1998), Phosphorus accumulation in marine sediments and the oceanic phosphorus cycle, Glob. Biogeochem. Cycles, 12(4), 563–572, doi:10.1029/98GB02263.

Ehrlich, H. L. (1998), Geomicrobiology: its significance for geology, Earth-Sci. Rev., 45(1), 45–60, doi:10.1016/S0012-8252(98)00034-8.

Hirschler, A., J. Lucas, and J.-C. Hubert (1990a), Apatite genesis: A biologically induced or biologically controlled mineral formation process?, Geomicrobiol. J., 8(1), 47–56, doi:10.1080/01490459009377877.

Hirschler, A., J. Lucas, and J.-C. Hubert (1990b), Bacterial involvement in apatite genesis, FEMS Microbiol. Ecol., 6(3), 211–220, doi:10.1111/j.1574- 6968.1990.tb03943.x.

Kallmeyer, J., R. Pockalny, R. R. Adhikari, D. C. Smith, and S. D’Hondt (2012), Global distribution of microbial abundance and biomass in subseafloor sediment, Proc. Natl. Acad. Sci., 109(40), 16213–16216, doi:10.1073/pnas.1203849109.

Krom, M. D., N. Kress, S. Brenner, and L. I. Gordon (1991), Phosphorus limitation of primary productivity in the eastern mediterranean sea, Limnol. Oceanogr., 36(3), 424–432, doi:10.4319/lo.1991.36.3.0424.

Parkes, R. J., B. A. Cragg, S. J. Bale, J. M. Getliff, K. Goodman, P. A. Rochelle, J. C. Fry, A. J. Weightman, and S. M. Harvey (1994), Deep bacterial biosphere in Pacific Ocean sediments, Lett. Nat., 371, 410–413, doi:10.1038/371410a0.

4 Roussel, E. G., M.-A. C. Bonavita, J. Querellou, B. A. Cragg, G. Webster, D. Prieur, and R. J. Parkes (2008), Extending the Sub-Sea-Floor Biosphere, Science, 320(5879), 1046–1046, doi:10.1126/science.1154545.

Ruttenberg, K. C. (1992), Development of a sequential extraction method for different forms of phosphorus in marine sediments, Limnol. Oceanogr., 37(7), 1460–1482, doi:10.4319/lo.1992.37.7.1460.

Ruttenberg, K. C., N. O. Ogawa, F. Tamburini, R. A. Briggs, N. D. Colasacco, and E. Joyce (2009), Improved, high-throughput approach for phosphorus speciation in natural sediments via the SEDEX sequential extraction method, Limnol. Oceanogr.-Methods, 7, 319–333, doi:10.4319/lom.2009.7.319.

Wu, J., W. Sunda, E. A. Boyle, and D. M. Karl (2000), Phosphate Depletion in the Western North Atlantic Ocean, Science, 289(5480), 759–762, doi:10.1126/science.289.5480.759.

Chapter One

A NEW SOLUTION 31P NMR SAMPLE PREPARATION SCHEME FOR

MARINE SEDIMENTS

Submitted: Defforey, D., Cade-Menun, B.J. and Paytan, A. (2016). A new solution 31P NMR sample preparation scheme for marine sediments. Limnol. Oceanogr. Methods, In Review.

6 Abstract

A new approach for the preparation of marine sediment samples for solution 31P nuclear magnetic resonance spectroscopy (31P NMR) has been developed and tested.

This approach addresses important aspects associated with sample pretreatments for marine sediments, including the effects of sample pretreatment on sedimentary P composition. The method increases the signals of low abundance P species in 31P

NMR spectra by quantitatively and precisely removing up to 80% of inorganic P

(orthophosphate) from sediment samples while causing minimal alteration of the chemical structure of organic P compounds. This method uses a reductive step to solubilize P bound to iron oxyhydroxides, followed by a low pH digestion to extract P from authigenic and biogenic apatite, as well as P bound to calcium carbonate. These

P forms combined represent the most abundant inorganic P reservoir in marine sediments. The sample residue is then extracted in an alkaline solvent, 0.25 M NaOH

31 with 0.05 M Na2EDTA, and processed for P NMR spectroscopy. The method was tested on natural marine sediment samples from different localities with high inorganic P content (> 85% molybdate reactive P), and allowed for the detection of orthophosphate monoesters and pyrophosphate in samples for which only an orthophosphate signal could be resolved with an NaOH-EDTA extraction alone. This new approach will allow the use of 31P NMR on samples for which low organic P concentrations previously hindered the use of this tool, and will help answer longstanding question regarding the fate of organic P in marine sediments.

7 1.1 Introduction

Phosphorus (P) is a critical macronutrient for life and thus plays a substantial role in regulating primary productivity in some marine systems and, over geological timescales, in the world’s oceans (Krom et al. 1991; Wu et al. 2000; Paytan and

McLaughlin 2007). In the open ocean, organic P derived from planktonic and bacterial biomass is the primary form in which P is delivered to marine sediments, and can make up 25 – 30% of total sedimentary P (Faul et al. 2005; Paytan and

McLaughlin 2007; Ruttenberg 2014). Despite its importance to marine P cycling, there is still a limited understanding of the chemical composition of organic P in sediments. This is largely due to low concentrations and analytical difficulties associated with isolating organic matter and determining the chemical speciation of organic P (Ruttenberg 2014). Most studies of marine P cycling in sediments have focused on quantifying total sedimentary organic P concentrations rather than identifying the compounds that make up the sedimentary P pool (Filippelli and

Delaney 1996; Anderson and Delaney 2001; Ruttenberg 2014). These studies have shown that the size of the bulk sedimentary organic P pool tends to decrease with depth, consistent with microbial respiration of organic P compounds, until it reaches an asymptotic value that remains low and at constant detectable values. However, mechanisms that would allow for the utilization of sedimentary organic P, the relative reactivity of distinct compounds within this pool and their degree of preservation during diagenesis are still poorly constrained. Numerous studies have proposed possible compounds within the organic P pool (inositol phosphates, phosphonates)

8 that are more refractory and could resist microbial respiration, only to reach the conclusion that these can be utilized for microbial respiration as readily as labile P compounds, such as phosphomonoesters and phosphodiesters (Suzumura and

Kamatani 1995; Laarkamp 2000; Benitez-Nelson et al. 2004). This implies that even organic P compounds previously thought to be refractory to microorganisms can be mineralized.

The application of solid-state and solution 31P nuclear magnetic resonance spectroscopy (hereafter referred to as 31P NMR) to marine sediments and sinking marine particulate matter has substantially improved our understanding of the composition of the bulk organic P sedimentary pool (Carman et al. 2000; Laarkamp

2000; Paytan et al. 2003; Sannigrahi and Ingall 2005). Indeed, 31P NMR studies of marine sediments from different localities indicate that orthophosphate typically constitutes 50 – 70% of total sedimentary P, pyrophosphate 0 – 4%, polyphosphate 0

– 8%, phosphonates 0 – 15%, P-monoesters ~30% and P-diesters up to 16% (Paytan et al. 2003; Sannigrahi and Ingall 2005). However, successful 31P NMR experiments require high organic to inorganic P ratios and high P concentrations, making 31P

NMR difficult to use with the more abundant open ocean low organic matter sediments. A small number of studies have used pretreatments to increase P concentrations and lower paramagnetic ion content in marine sediments. While these studies provided insightful results, some of them had limitations or lacked sufficient information to be reproduced. Carman et al. (2000) used a two-step pre-extraction

9 (citrate-dithionite-bicarbonate and magnesium chloride) followed by two double- distilled water rinses and a 24 h alkaline extraction (0.5 M NaOH) to prepare samples for solution 31P NMR. The 31P NMR experiments consisted of 30,000 scans (24 h); this, along with the duration of the alkaline extraction, raises the question of possible organic P hydrolysis during sample preparation and analysis. Ingall et al. (1990) and

Sannigrahi and Ingall (2005) used an HF/HCl pretreatment to remove mineral phases and concentrate organic matter. However, no information regarding the amount of P removed during pretreatment and the effects of the pretreatment on sample P compounds was provided. Since those studies were published, HF pretreatments have been shown to completely remove inositol phosphates and pyrophosphate from soil samples (Dougherty et al. 2007; Hamdan et al. 2012). This highlights the need for a detailed extraction procedure to prepare marine sediments for solution 31P NMR that includes information about the effects of the sample pretreatment on existing compounds and an evaluation of result reproducibility.

This study focuses on solution 31P NMR because it allows for sample pre- concentration and offers higher spectral resolution than solid-state NMR, and thus has the potential to identify more compounds (Cade-Menun et al., 2005). Since the output of 31P NMR experiments comes in the form of percentage of total sample P for each P species based on resolvable peak area, samples with high inorganic P content (>90%) can yield spectra that are dominated by the orthophosphate peak. While other P compounds may be present, their smaller peaks may not be sufficiently resolved from

10 the background noise; as such, the number of collected scans may have to be substantially increased to improve the signal-to-noise ratio enough to see them. The goal of this procedure is to amplify the signal of organic P compounds relative to that of orthophosphate so that low abundance P species may be resolved with 31P NMR without the need for extremely long 31P NMR experiments. Our sample preparation scheme solubilizes orthophosphate from mineral phases during the first two steps of the procedure before the sediment sample is extracted for solution 31P NMR. First, P bound to iron (Fe) oxyhydroxides is released from sediment samples through a reductive step during which sodium dithionite reduces Fe oxyhydroxides, and citrate complexes with Fe, thus releasing P associated with Fe oxides (Ruttenberg 1992).

Following this step, the sediment residue is extracted in a low pH sodium acetate buffer to dissolve authigenic, biogenic apatite, as well as calcium carbonate

(Ruttenberg 1992). We chose to use a sodium acetate buffer instead of a stronger acid for this step to minimize the alteration of the chemical structure of organic P compounds during the extraction. This two-step pretreatment quantitatively removes the majority of mineral P from marine sediment samples while also removing polyvalent bridging cations that can interfere with the alkaline extraction and speciation of organic P in marine sediments using 31P NMR (Turner et al. 2005). With this pretreatment, P species with low relative abundance that would not be detected using a single alkaline extraction can be quantified. Our method offers an accessible and reproducible means of preparing samples that would otherwise be unsuitable for

31P NMR analysis.

1.2 Materials and procedures

1.2.1 Materials

Laboratory supplies –We used 250 mL high-density polyethylene (HDPE) Nalgene bottles and 2 L HDPE bottles for reagent solutions. All glassware and plasticware were soaked in Decon Neutrad phosphate-free soap, acid-washed in 10% hydrochloric acid and rinsed with 18.2 MΩ·cm deionized water (hereafter referred to as ultrapure water) prior to usage.

Chemical supplies –All chemical supplies used for this study were ACS-grade. These include trisodium citrate (Fisher Chemical), sodium bicarbonate (Fisher Chemical), sodium dithionite (Acros Organics), sodium acetate (Fisher Chemical), sodium hydroxide (Fisher Chemical), disodium ethylenediaminetetraacetic acid (Fisher

Chemical), 17.4 M glacial acetic acid (Fisher Chemical) and 18.0 M sulfuric acid

(Fisher Chemical). Ancillary information regarding reagent solution preparation is provided in Appendix 1.

Equipment –We used a table-top centrifuge capable of 3,000 to 4,000 rpm [Thermo

Scientific IEC CL40], a laboratory furnace [Barnstead Thermolyne 62700], an analytical balance [Ohaus Explorer], a shaker table [New Brunswick Scientific C1

Platform Shaker], a freeze-drier [Labconco 75035], an inductively coupled plasma optical emission spectrometer (ICP-OES) [Perkin-Elmer Optima 4300 DV

12 Inductively Coupled Plasma Optical Emission Spectrometer operated by the

University of California, Santa Cruz], a QuikChem 8000 automated ion analyzer

[Lachat Instruments] and an NMR spectrometer [600 MHz Varian Unity INOVA spectrometer equipped with a 10 mm broadband probe].

1.2.2 Extraction procedure

The pretreatment (Fig. 1-1) used a reductive step to solubilize reducible P bound to

Fe oxyhydroxides (Pred), followed by a mild acid digestion to extract P from authigenic and biogenic apatite, as well as P bound to calcium carbonate (PCFA).

These P pools combined represent the most abundant inorganic P reservoirs in marine sediments (Ruttenberg, 1992). The residue was then extracted in an alkaline solvent,

0.25 M NaOH with 0.05 M Na2EDTA (hereafter referred to as NaOH-EDTA), which has been shown to be the most effective extractant solution for organic P in soils

(Cade-Menun and Preston 1996; Turner et al. 2005).

Initial sample P characterization– we used the ignition method to determine total P and molybdate-reactive P concentrations (MRP, which includes primarily free orthophosphate) for each sediment sample used for this study. Samples for total P analyses were ashed in crucibles at 550oC for 2 h and then extracted in 25 mL of 0.5

M sulfuric acid for 16 h. Samples for MRP analyses were extracted in the same manner, without the ashing step (Olsen and Sommers 1982; Cade-Menun and

Lavkulich 1997). We derived molybdate-unreactive P concentrations (MUP, which

13 includes primarily organic P and polyphosphates) by subtracting MRP from total P concentrations. For ashed and unashed extracts, MRP was determined as described below in the section titled “Supernatant P analysis”.

Sample pretreatment– Prior to the extraction, we freeze-dried, ground and sieved sediment samples to less than 125 µm (Ruttenberg 1992). For a given sample, we weighed four sample replicates (2 g) and placed each in 250 mL HDPE bottles.

Sodium dithionite (F.W. 147.12 g/mol; 7.4 g) was added to each sample split, followed by 200 mL of citrate-bicarbonate solution (pH 7.6). This step produces effervescence, so the solution should be added slowly to the sample. We shook samples for 8 h and then centrifuged them at 3,700 rpm for 15 min. We filtered the supernatants with a 0.4 µm polycarbonate filter. We took 20 mL aliquots from the filtrate for each sample split for MRP and total P analyses, and kept them refrigerated until analysis within 24 h. We added 200 mL of ultrapure water to the solid residue for each sample split as a wash step after the above reductive step, shook samples for

2 h, and then centrifuged them at 3,700 rpm for 15 min. We filtered the supernatants with 0.4 µm polycarbonate filters and set aside 20 mL of filtrate from each sample split for MRP and total P analyses. We then extracted the solid sample residues in 200 mL of sodium acetate buffer (pH 4.0) for 6 h. At the end of this extraction step, we centrifuged the bottles at 3,700 rpm for 15 min, filtered the supernatants with 0.4 µm polycarbonate filters and took a 20 mL aliquot of filtrate from each sample split for

MRP and total P analyses. We added 200 mL of ultrapure water to the solid residue

14 for each sample split as a wash step, shook samples for 2 h, and then centrifuged them at 3,700 rpm for 15 min. We filtered the supernatants with 0.4 µm polycarbonate filters and set aside 20 mL of filtrate from each sample split for MRP and total P analyses. We repeated the water rinse step, and collected aliquots for MRP and total P analyses as in the previous steps. The concentrations of total P and MRP were determined as described below in the section titled “Supernatant P analysis”.

Solution 31P NMR extraction– Solid sediment sample residues following the pretreatment described above were transferred to two 50 mL centrifuge tubes (2 sample replicates combined per tube). We added 20 mL of 0.25 M NaOH + 0.05 M

Na2EDTA solution to each tube, vortexed until all sediment was resuspended and then shook samples for 6 h at room temperature (Cade-Menun et al. 2005). We used a solid to solution ratio of 1:5 for this step to minimize the amount of freeze-dried material that will need to be dissolved for the 31P NMR experiments. Large amounts of salts from the NaOH-EDTA concentrated in NMR samples lead to higher viscosity and increase line broadening on NMR spectra (Cade-Menun and Liu 2014). We chose an extraction time of 6 h to improve total P recovery while limiting the degradation of natural P compounds in the sample. At the end of the extraction, samples were centrifuged at 3,700 rpm for 15 min and supernatants decanted into 50 mL centrifuge tubes. We collected a 500 µL aliquot from each sample, which we diluted with 4.5 mL of ultrapure water . These were refrigerated until analysis for total P content on the ICP-OES. The sample residues and supernatants were frozen on a slant to

15 maximize the exposed surface area during the lyophilization step; this was done immediately after the removal of the 500 µL aliquot. Once completely frozen, the uncapped tubes containing supernatants and residues were freeze-dried over the course of 48 h. Each tube was covered with parafilm with small holes from a tack to minimize contamination. Freeze-dried supernatants from identical sample splits were combined and dissolved in 500 µL each of ultrapure water, D2O, NaOH-EDTA and

31 10 M NaOH prior to P NMR analysis. The D2O is required as signal lock in the spectrometer (Cade-Menun and Liu 2014). Sample pH was maintained at a pH > 12 to optimize peak separation (Cade-Menun 2005; Cade-Menun and Liu 2014). Sample pH was assessed with a glass electrode, and verified with pH paper to account for the alkaline error caused by the high salt content of our samples (Covington 1985).

Solution 31P NMR analysis– Spectra were acquired immediately following sample preparation on a Varian Unity INOVA 600 MHz spectrometer equipped with a 10- mm broadband probe [operated by the Stanford Magnetic Resonance Laboratory at

Stanford University]. We used a 10-mm rather than a 5-mm probe because larger tubes contain a greater concentration of P and thus require fewer scans to achieve similar signal to noise ratios (Cade-Menun and Liu 2014). The analytical parameters used were: 20oC, 90o pulse, 0.48 s acquisition time, 4.52 s delay time, 5600 scans (8 h experiments), no spin and an external H3PO4 standard. We maintained samples at a temperature of 20oC during experiments to achieve optimal spectral resolution

(Crouse et al. 2000) and to minimize sample degradation. No proton decoupling was

16 used out of concern for sample degradation (Cade-Menun and Liu 2014). The ratio of

P to Fe and manganese (Mn) was used as a proxy for spin-lattice relaxation times (T1) to ensure adequate delays between pulses and thus quantitative spectra (McDowell et al. 2006). We used 5 s recycle delays, which correspond to three to five times the calculated T1 values, as recommended by McDowell et al. (2006). Peak identification was based on literature (Turner et al. 2003; Cade-Menun 2015).

Post-experimental processing– 31P NMR data were processed using the NMR Utility

Transform software (NUTS, Acorn NMR). Peak areas were calculated by integration of spectra processed with a 7 Hz line broadening following baseline correction, peak picking and phasing. We accepted peaks that (1) represented at least 1% of the tallest peak in the total integrated area, (2) were identified by the NUTS software and (3) were confirmed as signal by visual inspection.

Residual P extraction– Freeze-dried sample residues were ashed in crucibles at 550oC for 2 h and then extracted in 25 mL of 0.5 M sulfuric acid for 16 h (Olsen and

Sommers 1982; Cade-Menun and Lavkulich 1997). We centrifuged samples at 3,700 rpm for 15 min, filtered supernatants with 0.4 µm polycarbonate filters, and measured

P content on an ICP-OES.

Supernatant P analyses– Total P concentrations in sediment extracts were measured using inductively coupled plasma optical emission spectroscopy (ICP-OES).

17 Standards were prepared with the same solutions as those used for the extraction procedure in order to minimize matrix effects on P measurements. Sediment extracts and standards (0 µM, 3.2 µM, 32 µM and 320 µM) were diluted to lower salt content to prevent salt buildup on the nebulizer (1:20 dilution for step 1, 1:10 for steps 2 – 4).

Concentration data from both wavelengths (213 nm and 214 nm) were averaged to obtain extract concentrations for each sample. The detection limit for P on this instrument for both wavelengths is 0.4 µM. The MRP concentrations were measured on a QuikChem 8000 automated ion analyzer. Standards were prepared with the same solutions used for the extraction step to minimize matrix effects on P measurements.

Sediment extracts and standards (0 – 30 µM PO4) were diluted ten-fold to prevent matrix interference with color development. The detection limit for P on this instrument is 0.2 µM. We derived MUP concentrations by subtracting MRP from total P concentrations.

Degradation experiments– We verified the effects of our pretreatment on different classes of P compounds by spiking blank CDB and Na-acetate solutions with known standards, and monitoring the degree of hydrolysis over the course of the extraction period. Samples were analyzed by NMR for 13.3 min (16 scans) immediately after preparation of the spiked solutions; this was repeated every after 3 h (sodium acetate experiments) or 4 h (CDB experiments) of storage at room temperature. The standards used to assess hydrolysis due to sample pretreatment were 2- aminoethylphosphonic acid, D-glucose-6-phosphate, DL-α-glycerophosphate, RNA

18 and ATP. We used methylenedisphosphonic acid (MDP) as an internal standard in these experiments.

1.3 Assessment

1.3.1 Evaluation of extraction procedure on natural samples

We evaluated the performance of our method by testing it on natural marine sediment samples and comparing results to those obtained using the commonly used NaOH-

EDTA extraction without pretreatment. The sediment samples used were collected during USGS cruises P-1-94-AR (Arctic Ocean) and W-2-98-NC (California Margin) and the JGOFS cruise TT013-06MC (Equatorial Pacific). Samples were selected based on their latitude, differing sediment types and sedimentary P characteristics

(Table 1-1, Fig. 1-2). We chose samples with high MRP content (> 85% of total P), and a range of sedimentary mineral composition to assess the performance of our method (Table 1-1, Fig. 1-2). For the Arctic and Equatorial Pacific samples, our method allowed for the detection of P monoesters and pyrophosphate, which would otherwise not be detectable using a single NaOH-EDTA extraction (Table 1-2, Fig. 1-

3). For the samples near the California Margin, orthophosphate and P monoesters were detected using both the single NaOH-EDTA extraction and our method (Table

1-2, Fig. 1-4). However, our method allowed for the detection of pyrophosphate, which was not measurable with a single alkaline extraction. The total P monoester concentrations were similar when comparing both methods, showing that the pretreatment did not remove any measurable P monoesters (Table 1-2). We attempted

19 to identify specific monoester compounds using spiking, but this was not successful due to low signal to noise ratios and line broadening in the spectra. However, there appear to be some diester degradation peaks (α- and β-glycerophosphate) of similar magnitude in samples treated with the single NaOH-EDTA extraction and in the samples processed with the pretreatment we employed.

Extraction yields for samples with no pretreatment ranged between 1.8 – 23.6% of total P (Table 1-2). In comparison, extraction yields for samples with the pretreatment ranged between 78.6 – 86.3% (Table 1-2). The P content in the NMR extracts was lower in pretreated samples than in samples with no pretreatment because the pretreatment removed up to 80% of the samples’ MRP content (Table 1-

3, Fig. 1-5). However, adding the pretreatment allowed the detection of P compounds previously undetectable in the P NMR spectra, and increased overall extraction yields

(Table 1-2).

The pretreatment also resulted in lower concentrations of Al, Ca, K and Mg in the

NaOH-EDTA extracts (Table 1-3). However, it released additional paramagnetic ions

(Fe, Mn) compared to samples treated with a single NaOH-EDTA extraction, causing a decrease in spin-lattice relaxation times (Table 1-3). Without the pretreatment, the recycle delays in the NMR experiments used were too short to yield quantitative spectra for the Arctic Ocean samples and the Equatorial Pacific samples (Tables 1-2,

1-3). The higher paramagnetic ion content of the California Margin samples yielded

20 quantitative spectra even without the pretreatment since the recycle delays used were

3 to 5 times larger than calculated T1 times (Tables 1-2, 1-3). This supports the argument that the signal in the spectra for the Arctic Ocean samples and the

Equatorial Pacific samples was saturated with too short delay times.

It is difficult to conduct replicate analyses of all samples with 31P NMR due to the long experimental times involved and thus the expensive nature of these analyses. For these reasons, we determined the method’s reproducibility by extracting multiple replicates of only one of the samples (TT013-06MC 10-12 cm). Measurement uncertainties for MRP for the pretreatment steps ranged from 2.3 – 2.6% (steps 2 and

1 respectively, results not shown). Total P measurement uncertainties for each extraction step are reported in Table 1-4. Uncertainties ranged between 1.0 – 3.8% for the pretreatment steps, and increased to 8 – 12% for orthophosphate and monoester P determined via P NMR and for the residual P. The uncertainty associated with measuring pyrophosphate using P NMR is greater (28%) due to the low concentrations present in the samples (< 1% of total P), but similar to other published values (Xu et al. 2012).

1.3.2 Pretreatment effects on sample chemistry

We monitored the effects of the sample pretreatment on removal of P compounds other than MRP by measuring the MRP and total P content of each extraction step prior to the 31P NMR extraction. We derived MUP content in pretreatment extracts by

21 subtracting MRP from total P concentrations. For all samples, pretreatment extracts contained no detectable levels of MUP (Fig. 1-5), which indicates that our method does not remove or hydrolyze organic P prior to the 31P NMR extraction step.

We further assessed the effects of the sample pretreatment by monitoring the degree of hydrolysis of eight P compounds representative of naturally occurring organic P.

We spiked CDB and sodium acetate blanks with standards, and observed their relative concentrations over the course of the extraction period (8 h for CDB standards, 6 h for sodium acetate standards). We observed no degradation of D- glucose-6-phosphate and DL-α-glycerophosphate during the 8 h CDB extraction (Fig.

1-6). However, the CDB extraction altered the 2-aminoethylphosphonic acid signal and caused some hydrolysis over the course of 8 h (Fig. 1-6). A small decrease in

RNA and ATP was also observed after 8 h (Fig. 1-6). We did not observed degradation in 2-aminoethylphosphonic acid, D-glucose-6-phosphate and DL-α- glycerophosphate over the course of the 6 h sodium acetate extraction (Fig. 1-7).

However, there was a slight decrease in the abundance of RNA and ATP. Overall, the majority of the compounds tested were not rapidly degraded during the pretreatment.

It is important to note, however, that RNA, and to a lesser extent ATP, hydrolysis may occur over the course of sample preparation and should be taken into account when interpreting spectra. RNA has been shown to be susceptible to hydrolysis under alkaline conditions during NMR experiments (Smernik et al. 2015), so some degree of degradation with solution 31P NMR using NaOH-EDTA can be expected, with or

22 without any pre-treatment. However, quantification of hydrolysis products can help circumvent the loss of information due to RNA hydrolysis.

1.4 Discussion

The new method described here allows for the detection of P forms in marine sediments that were previously below detection limit by effectively removing up to

80% of mineral P and removing polyvalent bridging cations that can interfere with the alkaline extraction and speciation of organic P in marine sediments, thus increasing the relative signal of low abundance P species (P monoesters, pyrophosphate) in 31P NMR spectra. The reductive step solubilizes orthophosphate bound to Fe oxyhydroxides while removing no measurable MUP, causing no degradation to D-glucose-6-phosphate and DL-α-glycerophosphate, and minimal degradation to 2-aminoethylphosphonic acid, RNA and ATP. The sodium acetate extraction dissolves authigenic, biogenic apatite, as well as calcium carbonate and extracted no measurable levels of MUP. It is worth noting that Cade-Menun et al.

(2015) recently showed that sodium acetate can be used to extract P for NMR analyses in manure samples similarly to NaOH-EDTA. A possible explanation for our different results is that organic P forms are more stabilized and immobile in open ocean sediments than in manure, which affects the degree to which they can be solubilized in the sodium acetate extraction. We recommend future studies monitor the MRP and MUP content of supernatants during extraction to ensure that no significant amount of MUP is solubilized. The sodium acetate extraction did not

23 cause any observable degradation in 2-aminoethylphosphonic acid, D-glucose-6- phosphate and DL-α-glycerophosphate. However, some degradation of RNA and

ATP was observed and could result in underestimations of these compounds.

Without this new pretreatment, spectra from the Equatorial Pacific and Arctic sediment samples were dominated by orthophosphate, leaving no other peaks visible

(Table 1-2, Fig. 1-3). The organic P compounds measured in our samples were orthophosphate monoesters (Table 1-2, Fig. 1-3, 1-4). As mentioned above, we were unable to resolve individual peaks in the monoester region due to line broadening and low signal to noise ratios. While identifying peaks in the monoester region was beyond the scope of this methods paper, we recommend future studies correct for degradation peaks to accurately calculate the total amount of orthophosphate monoesters and diesters. Orthophosphate monoesters are generally the dominant group of organic P compounds in both marine and terrestrial environments, and include mononucleotides, sugar phosphates, inositol phosphates and phosphoproteins

(Ingall et al. 1990; Clark et al. 1998, 1999; Carman et al. 2000; Kolowith et al. 2001;

Turner et al. 2005; Sannigrahi and Ingall 2005; Sannigrahi et al. 2006). Certain orthophosphate monoesters, particularly the inositol hexakisphosphate stereoisomers, have a high affinity for clays, Fe and aluminum oxide and oxyhydroxide surfaces that leads to the formation of insoluble complexes with polyvalent cations (Celi and

Barberis 2005). These complexes may increase their resistance to microbial degradation and lead to their accumulation in sediments (Celi and Barberis 2005).

24 Orthophosphate monoesters are thought to adsorb on the same sites as phosphate anions by exchanging ligands with the mineral surface hydroxyl groups (Baldwin et al. 2002; Celi and Barberis 2005). Orthophosphate diesters, on the other hand, adsorb more weakly on mineral surfaces and are more readily mineralizable, which makes them more vulnerable to microbial degradation (Paytan et al. 2003; Celi and Barberis

2005). Furthermore, organic P remineralization occurs throughout the water column in addition to sediments (Paytan et al. 2003; Benitez-Nelson et al. 2007). These mechanisms combined could account for the absence of orthophosphate diesters in our spectra. Another possibility would be that some orthophosphate diesters were hydrolyzed to orthophosphate monoesters during the extraction process or during

NMR analysis (Schneider et al., 2016). Resolving individual monoester peaks in future studies will allow the identification of diester degradation peaks (α- and β- glycerophosphate, mononucleotides) and enable corrections to accurately calculate the total amount of orthophosphate monoesters and diesters separately (Schneider et al. 2016).

We found that all six sediment samples contained small amounts of pyrophosphate, a biogenic form of inorganic P (Table 1-2). Indeed, pyrophosphate is a product of the biosynthesis of a number of macromolecules, including nucleic acids, amino acids, coenzymes, nucleotides and activated precursors to proteins and polysaccharides

(Heinonen 2001). Pyrophosphate has been measured in a number of natural samples, including plankton, sediment trap material and sediments (Sundareshwar et al. 2001;

25 Paytan et al. 2003; Cade-Menun et al. 2005). As in Paytan et al. (2003), sedimentary pyrophosphate concentrations are low and decrease slightly with increasing latitude, supporting the hypothesis of surface water temperature possibly influencing the rate of synthesis or decomposition of this compound.

It is interesting to note that phosphonates, which contain carbon-P bonds, were not detected in any of our sediment samples. Phosphonates have been measured in many marine samples, including sediment trap material, sediments, and dissolved and particulate organic matter (Ingall et al. 1990; Clark et al. 1998, 1999; Carman et al.

2000; Kolowith et al. 2001; Paytan et al. 2003; Benitez-Nelson et al. 2004; Cade-

Menun et al. 2005; Sannigrahi et al. 2006). There are two possible hypotheses that could account for the lack of phosphonate detection in our spectra:

(1) Phosphonates were remineralized in the water column as particulates sank and in surface sediments, causing sedimentary phosphonates to be either absent or below detection limit despite the pretreatment. Paytan et al. (2003) reported that phosphonates represented 1% of sedimentary P at two sites in the Equatorial Pacific.

We did not observe phosphonates in our Equatorial Pacific samples; however it is important to note that our samples were taken from greater depths (10 – 12 cmbsf and

24 – 26 cmbsf compared to 1 – 3 cmbsf in Paytan et al. 2003). It is possible that phosphonates were remineralized in the overlying sediments, causing their signal to decrease with depth. Indeed, phosphonate abundance has been shown to decrease significantly between the sediment-water interface and 2 cmbsf in anoxic sediments

26 relative to orthophosphate and P esters (Benitez-Nelson et al. 2004). Phosphonate remineralization upon sediment deposition could explain the absence of a phosphonate signal in our spectra.

(2) Another possible explanation for the absence of a phosphonate signal is that the alkaline extraction step did not completely solubilize phosphonates. Indeed, while extracting samples with NaOH-EDTA removes the majority of P esters, this alkaline extraction removes a variable portion of phosphonates from marine particulate samples (Cade-Menun et al. 2005). If sedimentary phosphonate levels are low, an incomplete solubilization will result in phosphonate signals below detection limit for

31P NMR spectroscopy. Adding an additional step in the procedure involving solid- state 31P NMR of sediment residues prior to ashing could provide information regarding any organic P compounds that were not extracted.

This new method enhances the use of solution 31P NMR as an effective tool to study organic P in sediments with low organic P content (less than 10% of total P), which represent a large fraction of marine sediments. The use of 31P NMR until now has offered valuable insights into P cycling in sediments with high organic P content, but was not as effective and easily implementable with sediments containing high inorganic to organic P ratios. Using the pre-extraction preparation step, we were able to detect and identify the chemical structure of low abundance organic P species in sediments and demonstrated that orthophosphate monoesters persist in marine sediments despite their potential bioavailability to deep sedimentary subseafloor

27 microorganisms. Furthermore, we observed that pyrophosphate is often present in marine sediment samples and that its abundance varies with latitude. It would be interesting to use this new method to monitor changes in organic P forms and abundance with depth in sediments to investigate the impact of microbial activity and diagenetic processes on organic P compounds as a function of burial depth and age.

This will provide valuable insights into the factors controlling the relative reactivity and lability of P compounds and the preservation of organic P in sediments.

1.5 Comments and recommendations

Chemical shifts are expressed relative to an external standard and are pH-dependent.

For this reason, it is important to consider the standard used and sample pH when comparing results from different studies (Cade-Menun 2015). Maintaining sample pH above 12 and temperature at 20oC during the NMR experiment is optimal for peak separation. We recommend using pH indicator strips in addition to a glass electrode to measure the pH of redissolved NaOH-EDTA extracts to avoid inaccuracies caused by their high salt content, which interferes with standard glass electrode measurements.

When possible, spiking with standards should be used to identify specific compounds in P NMR spectra in addition to comparing peaks with compound libraries from published literature. This is the most reliable way to confirm peak identifications.

However, given the cost associated with purchasing a large number of pure

28 compounds for spiking, peak libraries have been published and provide a cost- effective means of identifying peaks (Turner et al. 2003; Cade-Menun 2015).

Another important parameter to monitor when preparing marine sediments for 31P

NMR analysis is the paramagnetic ion content of the NaOH-EDTA extracts using

ICP-OES. Paramagnetic ions affect spectral resolution, as well as recycle delays so their abundance needs to be determined prior to carrying out an NMR experiment to ensure that spectra are quantitative.

Acknowledgments

We thank the party members from USGS cruises P-1-94-AR and W-2-98-NC and those from the JGOFS cruise for collecting the sediment samples used in this study.

We also thank Michael Torresan and Cathy Frazee for providing us with sediment samples from the USGS cruises, and for their assistance. Funding was provided by a graduate fellowship from the Center for Dark Energy Biosphere Investigations (NSF

C-DEBI) to D.D. (award # 157598) and a research grant from C-DEBI to A.P. (award

# 156246). We also thank Dr. Corey Liu, Rob Franks and Michael Kong for their assistance with this project.

29 References

Anderson, L. D., and M. L. Delaney. 2001. Carbon to phosphorus ratios in sediments:

implications for nutrient cycling. Glob. Biogeochem. Cycles 15: 65–79.

doi:10.1029/2000GB001270

Baldwin, D. S., A. M. Mitchell, and J. M. Olley. 2002. Pollutant-sediment

interactions: sorption, reactivity and transport of phosphorus, p. 265–280. In

P.M. Haygarth and S.C. Jarvis [eds.], Agriculture, Hydrology and Water

Quality. CABI Publ.

Benitez-Nelson, C. R., L. O’Neill, L. C. Kolowith, P. Pellechia, and R. Thunell. 2004.

Phosphonates and particulate organic phosphorus cycling in an anoxic marine

basin. Limnol. Oceanogr. 49: 1593–1604. doi:10.4319/lo.2004.49.5.1593

Benitez-Nelson, C. R., L. P. O’Neill Madden, R. M. Styles, R. C. Thunell, and Y.

Astor. 2007. Inorganic and organic sinking particulate phosphorus fluxes

across the oxic/anoxic water column of Cariaco Basin, Venezuela. Mar.

Chem. 105: 90–100. doi:10.1016/j.marchem.2007.01.007

Cade-Menun, B. J. 2005. Characterizing phosphorus in environmental and

agricultural samples by 31P nuclear magnetic resonance spectroscopy. Talanta

66: 359–371. doi:10.1016/j.talanta.2004.12.024

Cade-Menun, B. J. 2015. Improved peak identification in 31P-NMR spectra of

environmental samples with a standardized method and peak library.

Geoderma 257-258: 102–114. doi:10.1016/j.geoderma.2014.12.016

30 Cade-Menun, B. J., C. R. Benitez-Nelson, P. Pellechia, and A. Paytan. 2005. Refining

31P nuclear magnetic resonance spectroscopy for marine particulate samples:

storage conditions and extraction recovery. Mar. Chem. 97: 293–306.

doi:10.1016/j.marchem.2005.05.005

Cade-Menun, B. J., Z. He, and Z. Dou. 2015. Comparison of phosphorus forms in

three extracts of dairy feces by solution 31P NMR analysis. Commun. Soil

Sci. Plant Anal. 46: 1698–1712. doi:10.1080/00103624.2015.1047512

Cade-Menun, B. J., and L. M. Lavkulich. 1997. A comparison of methods to

determine total, organic, and available phosphorus in forest soils. Commun.

Soil Sci. Plant Anal. 28: 651–663. doi:10.1080/00103629709369818

Cade-Menun, B. J., and C. W. Liu. 2014. Solution phosphorus-31 nuclear magnetic

resonance spectroscopy of soils from 2005 to 2013: a review of sample

preparation and experimental parameters. Soil Sci. Soc. Am. J. 78: 19.

doi:10.2136/sssaj2013.05.0187dgs

Cade-Menun, B. J., and C. M. Preston. 1996. A comparison of soil extraction

procedures for 31P NMR spectroscopy. Soil Sci. 161: 770–785.

doi:10.1097/00010694-199611000-00006

Carman, R., G. Edlund, and C. Damberg. 2000. Distribution of organic and inorganic

phosphorus compounds in marine and lacustrine sediments: a 31P NMR

study. Chem. Geol. 163: 101–114. doi:10.1016/S0009-2541(99)00098-4

31 Celi, L., and E. Barberis. 2005. Abiotic stabilization of organic phosphorus in the

environment, p. 113–132. In B.L. Turner, E. Frossard, and D.S. Baldwin

[eds.], Organic phosphorus in the environment. CABI Publ.

Clark, L. L., E. D. Ingall, and R. Benner. 1998. Marine phosphorus is selectively

remineralized. Nature 393: 426–426. doi:10.1038/30881

Clark, L. L., E. D. Ingall, and R. Benner. 1999. Marine organic phosphorus cycling;

novel insights from nuclear magnetic resonance. Am. J. Sci. 299: 724–737.

doi:10.2475/ajs.299.7-9.724

Covington, A. K. 1985. Procedures for testing pH responsive glass electrodes at 25,

37, 65 and 85oC and determination of alkaline errors up to 1 mol dm-3 Na+,

K+, Li+. Pure Appl Chem 57: 887–898. doi:10.1351/pac198557060887

Crouse, D. A., H. Sierzputowska‐Gracz, and R. L. Mikkelsen. 2000. Optimization of

sample pH and temperature for phosphorus‐31 nuclear magnetic resonance

spectroscopy of poultry manure extracts. Commun. Soil Sci. Plant Anal. 31:

229–240. doi:10.1080/00103620009370432

Dougherty, W. J., R. J. Smernik, E. K. Bünemann, and D. J. Chittleborough. 2007.

On the use of hydrofluoric acid pretreatment of soils for phosphorus-31

nuclear magnetic resonance analyses. Soil Sci. Soc. Am. J. 71: 1111.

doi:10.2136/sssaj2006.0300

Faul, K. L., A. Paytan, and M. L. Delaney. 2005. Phosphorus distribution in sinking

oceanic particulate matter. Mar. Chem. 97: 307–333.

doi:10.1016/j.marchem.2005.04.002

32 Filippelli, G. M., and M. L. Delaney. 1996. Phosphorus geochemistry of equatorial

Pacific sediments. Geochim. Cosmochim. Acta 60: 1479–1495.

doi:10.1016/0016-7037(96)00042-7

Griggs, G. B., and J. R. Hein. 1980. Sources, dispersal, and clay mineral composition

of fine-grained sediment off central and northern California. J. Geol. 541–566.

doi:10.1086/628543

Hamdan, R., H. M. El-Rifai, A. W. Cheesman, B. L. Turner, K. R. Reddy, and W. T.

Cooper. 2012. Linking phosphorus sequestration to carbon humification in

wetland soils by 31P and 13C NMR spectroscopy. Environ. Sci. Technol. 46:

4775–4782. doi:10.1021/es204072k

Heinonen, J. 2001. Biological role of inorganic pyrophosphate, Kluwer Academic

Publishers.

Ingall, E. D., P. A. Schroeder, and R. A. Berner. 1990. The nature of organic

phosphorus in marine sediments: new insights from 31P NMR. Geochim.

Cosmochim. Acta 54: 2617–2620. doi:10.1016/0016-7037(90)90248-J

Kolowith, L. C., E. D. Ingall, and R. Benner. 2001. Composition and cycling of

marine organic phosphorus. Limnol. Oceanogr. 46: 309–320.

doi:10.4319/lo.2001.46.2.0309

Krom, M. D., N. Kress, S. Brenner, and L. I. Gordon. 1991. Phosphorus limitation of

primary productivity in the eastern mediterranean sea. Limnol. Oceanogr. 36:

424–432. doi:10.4319/lo.1991.36.3.0424

33 Laarkamp, K. L. 2000. Organic phosphorus in marine sediments: chemical structure,

diagenetic alteration, and mechanisms of preservation. PhD Thesis. MIT-

WHOI Joint Program.

McDowell, R. W., I. Stewart, and B. J. Cade-Menun. 2006. An examination of spin–

lattice relaxation times for analysis of soil and manure extracts by liquid state

phosphorus-31 nuclear magnetic resonance spectroscopy. J. Environ. Qual.

35: 293. doi:10.2134/jeq2005.0285

Murray, R. W., and M. Leinen. 1993. Chemical transport to the seafloor of the

equatorial Pacific Ocean across a latitudinal transect at 135oW: tracking

sedimentary major, trace, and rare earth element fluxes at the Equator and the

Intertropical Convergence Zone. Geochim. Cosmochim. Acta 57: 4141–4163.

doi:10.1016/0016-7037(93)90312-K

Olsen, S. R., and L. E. Sommers. 1982. Phosphorus, p. 403–430. In A.L. Page, R.H.

Miller, and D.R. Keeney [eds.], Methods of Soil Analysis. Soil Science

Society of America.

Paytan, A., B. J. Cade-Menun, K. McLaughlin, and K. L. Faul. 2003. Selective

phosphorus regeneration of sinking marine particles: evidence from 31P-

NMR. Mar. Chem. 82: 55–70. doi:10.1016/S0304-4203(03)00052-5

Paytan, A., and K. McLaughlin. 2007. The Oceanic Phosphorus Cycle. Chem. Rev.

107: 563–576. doi:10.1021/cr0503613

34 Ruttenberg, K. C. 1992. Development of a sequential extraction method for different

forms of phosphorus in marine sediments. Limnol. Oceanogr. 37: 1460–1482.

doi:10.4319/lo.1992.37.7.1460

Ruttenberg, K. C. 2014. The Global Phosphorus Cycle, p. 499–558. In H. Holland

and K. Turekian [eds.], Treatise on Geochemistry. Elsevier.

Sannigrahi, P., and E. Ingall. 2005. Polyphosphates as a source of enhanced P fluxes

in marine sediments overlain by anoxic waters: evidence from P-31 NMR.

Geochem Trans 6: 52–59. doi:10.1063/1.1946447

Sannigrahi, P., E. D. Ingall, and R. Benner. 2006. Nature and dynamics of

phosphorus-containing components of marine dissolved and particulate

organic matter. Geochim. Cosmochim. Acta 70: 5868–5882.

doi:10.1016/j.gca.2006.08.037

Schneider, K. D., B. J. Cade-Menun, D. H. Lynch, and R. P. Voroney. 2016. Soil

phosphorus forms from organic and conventional forage fields. Soil Sci. Soc.

Am. J. 80: 328. doi:10.2136/sssaj2015.09.0340

Smernik, R. J., A. L. Doolette, and S. R. Noack. 2015. Identification of RNA

hydrolysis products in NaOH-EDTA extracts using 31P NMR spectroscopy.

Commun. Soil Sci. Plant Anal. 46: 2746–2756.

doi:10.1080/00103624.2015.1093640

Stein, R., H. Grobe, and M. Wahsner. 1994. Organic carbon, carbonate, and clay

mineral distributions in eastern central Arctic Ocean surface sediments. Mar.

Geol. 119: 269–285. doi:10.1016/0025-3227(94)90185-6

35 Sundareshwar, P. V., J. T. Morris, P. J. Pellechia, H. J. Cohen, D. E. Porter, and B. C.

Jones. 2001. Occurrence and ecological implications of pyrophosphate in

estuaries. Limnol. Oceanogr. 46: 1570–1577. doi:10.4319/lo.2001.46.6.1570

Suzumura, M., and A. Kamatani. 1995. Mineralization of inositol hexaphosphate in

aerobic and anaerobic marine sediments: implications for the phosphorus

cycle. Geochim. Cosmochim. Acta 59: 1021–1026. doi:10.1016/0016-

7037(95)00006-2

Turner, B. L., B. J. Cade-Menun, L. M. Condron, and S. Newman. 2005. Extraction

of soil organic phosphorus. Talanta 66: 294–306.

doi:10.1016/j.talanta.2004.11.012

Turner, B. L., N. Mahieu, and L. M. Condron. 2003. Phosphorus-31 nuclear magnetic

resonance spectral assignments of phosphorus compounds in soil NaOH-

EDTA extracts. Soil Sci Soc Am J 67: 497–510. doi:10.1016/S0146-

6380(03)00061-5

Wu, J., W. Sunda, E. A. Boyle, and D. M. Karl. 2000. Phosphate Depletion in the

Western North Atlantic Ocean. Science 289: 759–762.

doi:10.1126/science.289.5480.759

Xu, D., S. Ding, B. Li, F. Jia, X. He, and C. Zhang. 2012. Characterization and

optimization of the preparation procedure for solution P-31 NMR analysis of

organic phosphorus in sediments. J. Soils Sediments 12: 909–920.

doi:10.1007/s11368-012-0510-4

36 Step 1 Sample Iron-Bound P CDB (8 h)

Residue

H2O (2 h)

Step 2 Residue Auth. / bio. apatite Na acetate CaCO -bound P (6 h) 3 Residue

H2O (2 h) × 2

Step 3 Residue Base-extractable P NaOH-EDTA (NMR extract) (6 h) Step 4 Residue Residual P Ashing (2 h) H SO (16 h) 2 4 Fig. 1-1. Stepwise extraction scheme summarizing the preparation of marine sediment samples for solution 31P NMR spectroscopy.

37 a. b. 180 100

160 90

) 140 -1 70 120

mol g 60 μ 100 50 80 TT013-06MC TT013-06MC P-1-94-AR P21 P-1-94-AR P21 40 TF1 W-2-98-NC TF1 W-2-98-NC

phase P ( 60 − 30

40 Proportion of total P (%) Solid 20

20 10

0 0

0-4 cm 0-4 cm 16-20 cm 18-20 cm 24-26 cm P-1-94-AR P-1-94-AR W-2-98-NCW-2-98-NC 10-12 cm P21 0-4 cm TF1 0-4 cm TT013-06MC10-12 cmTT013-06MC24-26 cm P 21 16-20 cm TF1 18-20 cm TP MRP MUP MRP MUP

Fig. 1-2. (a) Total P, molybdate reactive P (MRP) and molybdate unreactive P (MUP) content of the sediment samples used for this study and (b) relative proportions of MRP and MUP for each sample. Total P concentrations were determined following the extraction of ignited sediment samples, while MRP concentrations were measured following the extraction of unignited sediment samples. All P concentration measurements were done colorimetrically on a QuikChem 8000 automated ion analyzer. MUP was determined by subtracting MRP from total P concentrations.

38 a. No pretreatmentTT013-06MC b. Pretreatment TT013-06MC 24-26 cm 24-26 cm

Orthophosphate Orthophosphate Orthophosphate monoester

Pyrophosphate

105 0 -5

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Fig. 1-3. (a) 31P NMR spectrum of sample TT013-06MC (24-26 cm) following a single NaOH-EDTA with no sample pretreatment and (b) spectrum of the same sample after pretreatment to remove orthophosphate. The inset spectrum in (b) is enlarged to better show details in the orthophosphate monoester and pyrophosphate regions. Spectra are plotted with 7-Hz line broadening, and are plotted with the orthophosphate peak at the same height.

39 a. No pretreatmentW-2-98-NC TF1 b. Pretreatment W-2-98-NC TF1 0-4 cm 0-4 cm

Orthophosphate Orthophosphate Orthophosphate monoester Orthophosphate monoester

R R

Pyrophosphate

105 0 105 0 -5

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Fig. 1-4. (a) 31P NMR spectrum of sample W-2-98-NC TF1 (0-4 cm) following a single NaOH-EDTA with no sample pretreatment and (b) spectrum of the same sample after pretreatment to remove orthophosphate. The inset spectra are enlarged to better show details in the orthophosphate monoester and pyrophosphate regions. Spectra are plotted with 7-Hz line broadening, and are plotted with the orthophosphate peak at the same height.

40 a. b. 35 100 ) )

-1 Step 1 -1 Step 2 30 90 80 mol g mol g 25 70 μ μ 20 60 50 15 40 phase P ( phase P ( 10 30 − − 20 5 Solid Solid 10 0 0

P-1-94-AR0-4 cmP-1-94-AR W-2-98-NCW-2-98-NC P-1-94-AR0-4 cmP-1-94-AR W-2-98-NCW-2-98-NC 16-20 cm 0-4 cm 16-20 cm 0-4 cm 18-20 TT013-06MCcm 10-12 cmTT013-06MC24-26 cm 18-20 TT013-06MCcm 10-12 cmTT013-06MC24-26 cm

MRP TP

Fig. 1-5. Mean MRP and total P content in pretreatment extracts from steps 1 (CDB and water extracts, panel a) and 2 (sodium acetate and water extracts, panel b), along with standard deviations derived from replicate sediment sample splits. Total P concentrations were measured on an ICP-OES, while MRP concentrations were measured colorimetrically on a QuikChem 8000 automated ion analyzer.

41 a. Extraction time: 0 h b. Extraction time: 4 h c. Extraction time: 8 h

OrthoP

MDP

AEP Gluc. PO4

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Chemical shift (ppm)

OrthoP

MDP RNA ATP

α-glyc.

PO4

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Chemical shift (ppm) Fig. 1-6. Spectra from the degradation test for the 8 h CDB extraction. Spectra in the top row are from a blank CDB solution spiked with 2-aminoethylphosphonic acid (AEP), methylenedisphosphonic acid (MDP), orthophosphate (orthoP) and D- glucose-6-phosphate (gluc. PO4). Spectra in the bottom row are from a blank CDB solution with with MDP, orthophosphate, DL-α-glycerophosphate (α –glyc. PO4), RNA and ATP. All solutions were pH adjusted to > 12. Spectra are plotted with 7-Hz line broadening, and are plotted with the orthophosphate peak at the same height.

42 a. Extraction time: 0 h b. Extraction time: 3 h c. Extraction time: 6 h

OrthoP MDP

AEP Gluc. PO4

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Chemical shift (ppm)

OrthoP

MDP RNA ATP

α-glyc.

PO4

30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 30 2010 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm) Chemical shift (ppm) Fig. 1-7. Spectra from the degradation test for the 6 h sodium acetate extraction. Spectra in the top row are from a blank sodium acetate solution spiked with 2- aminoethylphosphonic acid (AEP), methylenedisphosphonic acid (MDP), orthophosphate (orthoP) and D-glucose-6-phosphate (gluc. PO4). Spectra in the bottom row are from a blank sodium acetate solution with with MDP, orthophosphate, DL-α-glycerophosphate (α –glyc. PO4), RNA and ATP. All solutions were pH adjusted to > 12. Spectra are plotted with 7-Hz line broadening, and are plotted with the orthophosphate peak at the same height.

43 Table 1-1. Sample location, depth, sampling time and sedimentary characteristics of the samples used in this study

Water Sediment Sediment Sample ID Latitude, longitude Sampling dates depth (m) depth (cm) Composition

Arctic Ocean

P-1-94-AR P21 0-4 cm 84o5' N, 174o58' W 3,180 0 - 4 07/25/94 - 08/30/94 <10% CaCO3, high illite, low kaolinite P-1-94-AR P21 16-20 cm 84o5' N, 174o58' W 3,180 16 - 20 07/25/94 - 08/30/94 and chlorite1

California margin

W-2-98-NC TF1 0-4 cm 41o5' N, 125o1' W 3,045 0 - 4 07/17/98 - 07/24/98 High kaolinite and chlorite, moderate W-2-98 NC TF1 18-20 cm 41o5' N, 125o1' W 3,045 18 - 20 07/17/98 - 07/24/98 illite2

Equatorial Pacific

o o TT013-06MC 10-12 cm 12 00' S, 134 56' W 4,280 10 - 12 10/30/92 - 12/13/92 80% CaCO3, 10% opal3 TT013-06MC 24-26 cm 12o00' S, 134o56' W 4,280 24 - 26 10/30/92 - 12/13/92

1(Stein et al. 1994); 2(Griggs and Hein 1980); 3(Murray and Leinen 1993)

44 Table 1-2. Extraction yields for each step. Pred refers to P solubilized during the reductive step (step 1), PCFA refers to P released during the mild acid digestion (step 2). Orthophosphate (orthoP), orthophosphate monoester and pyrophosphate (pyroP) content was determined using 31P NMR (step 3). Extracted P is the fraction of P that was extracted in steps 1-3. Residual P (Res. P) is P remaining following the full extraction procedure.

———from 31P NMR spectra— —— NMR OrthoP P P OrthoP PyroP Res. P Extracted Sample ID red CFA extract TP monoesters (µmol/g) (µmol/g) (µmol/g) (µmol/g) (µmol/g) P (%) (µmol/g) (µmol/g)

Sample pretreatment

P-1-94-AR P21 0-4 cm 8.10 8.45 0.49 0.40 0.08 0.02 2.81 85.9 P-1-94-AR P21 16-20 cm 8.54 8.04 0.36 0.32 0.03 0.01 2.68 86.3 W-2-98-NC TF1 0-4 cm 4.89 11.9 1.56 1.36 0.16 0.03 4.04 82.0 W-2-98 NC TF1 18-20 cm 3.58 10.0 1.54 1.25 0.27 0.02 4.10 78.6 TTN013 10-12 cm 14.5 52.2 1.61 1.43 0.15 0.03 16.4 80.6 TTN013 24-26 cm 31.6 95.1 2.79 2.55 0.20 0.04 27.8 82.3

No pretreatment

P-1-94-AR P21 0-4 cm — — 2.31 — — — 14.0 14.2 P-1-94-AR P21 16-20 cm — — 3.93 — — — 12.8 23.6 W-2-98-NC TF1 0-4 cm — — 2.87 2.70 0.17 0.00 16.9 14.5 W-2-98 NC TF1 18-20 cm — — 1.97 1.66 0.31 0.00 14.8 11.8 TT013-06MC 10-12 cm — — 1.73 — — — 83.8 2.0 TT013-06MC 24-26 cm — — 2.53 — — — 141.6 1.8

45 Table 1-3. Aluminum, calcium, iron, potassium, magnesium, manganese and 31 phosphorus content in P NMR extracts. T1 values are derived from P to iron and manganese ratios (w/v) using the relationship derived by McDowell et al. (2006).

P Al Ca Fe K Mg Mn P Calculated Sample ID (Fe+Mn) T1 (s) ———————————µmol g-1———————————

Sample pretreatment P-1-94-AR P21 0-4 cm 1.31 12.3 0.26 0.28 4.54 0.02 0.49 1.00 0.60 P-1-94-AR P21 16-20 cm 1.70 13.2 0.35 0.25 4.97 0.02 0.36 0.56 0.47 W-2-98-NC TF1 0-4 cm 3.75 28.9 0.61 0.60 3.63 0.09 1.56 1.52 0.75 W-2-98 NC TF1 18-20 cm 2.88 17.3 0.63 0.62 2.68 0.05 1.54 1.28 0.68 TT013-06MC 10-12 cm 0.57 39.8 0.16 0.12 0.32 0.03 1.61 3.95 1.44 TT013-06MC 24-26 cm 0.46 51.4 0.43 0.24 0.36 0.05 2.79 3.23 1.24

No pretreatment P-1-94-AR P21 0-4 cm 5.17 58.1 0.09 11.4 35.0 0.00 2.31 16.7 5.09 P-1-94-AR P21 16-20 cm 3.76 120.0 0.06 12.3 43.1 0.01 3.93 38.7 11.4 W-2-98-NC TF1 0-4 cm 4.83 56.0 0.41 14.3 28.0 0.11 2.87 3.86 1.42 W-2-98 NC TF1 18-20 cm 4.84 54.2 0.52 15.0 34.5 0.06 1.97 2.36 0.99 1244. TT013-06MC 10-12 cm 3.06 0.00 82.3 0.06 0.02 1.73 105.1 30.4 3 1274. TT013-06MC 24-26 cm 1.99 0.00 103.8 0.08 0.03 2.53 111.7 32.3 1

46 Table 1-4. Measurement uncertainties associated with each extraction step using ICP- OES and P NMR spectroscopy determined by replicate analysis of sample TT013- 06MC (10-12 cm). Total P relative error is calculated as: relative error (%)= (SD/mean) × 100.

Mean TP Standard TP Rel. error Extraction step P fraction (µmol/g) deviation (%)

1 Pred 14.6 0.55 3.8 2 PCFA 52.7 0.54 1.0 OrthoP 1.34 0.11 7.9 3 MonoP 0.16 0.02 11.8 PyroP 0.03 0.01 27.9 4 Residual P 14.7 1.76 12.0

Chapter Two

PHOSPHORUS GEOCHEMISTRY OF SEDIMENTS IN RIDGE–FLANK

SYSTEMS

Published: Defforey, D. and Paytan, A. (2015). Data Report: Characteristics of sedimentary phosphorus at North Pond, IODP Expedition 336. In Edwards, K.J., Bach,W., Klaus, A., and the Expedition 336 Scientists, Proc. IODP, 336: Tokyo (Integrated Ocean Drilling Program Management International, Inc.). doi:10.2204/iodp.proc.336.205.2015

In prep: Defforey, D., LeVay, L. and Paytan, A. (2016). Phosphorus geochemistry in ridge-flank systems. Geochem Geophys. Geosys. (in prep)

Abstract

In this study, we present high-resolution solid-phase phosphorus (P) data from two well-studied sites to investigate the nature and processes altering sedimentary P in ridge-flank systems. While P in coastal marine sediments is well-studied thanks to the application of sequential extractions schemes, fewer datasets are available for P in open ocean sediment, particularly ridge-flank environments. The sediment samples used for this work were collected during Integrated Ocean Drilling Program (IODP)

Expedition 327 to the Juan de Fuca Ridge flank area, and IODP Expedition 336 to

North Pond, a sediment-filled basin located on the western flank of the Mid-Atlantic

Ridge. Solid-phase P reservoirs in sediments were characterized using the SEDEX sequential extraction scheme. This method quantitatively separates five distinct sedimentary P reservoirs based on each targeted phase’s reactivity: (1) loosely-sorbed

P; (2) ferric iron-bound P; (3) authigenic carbonate fluorapatite + biogenic apatite +

CaCO3-associated P; (4) detrital apatite and (5) organic P. In addition, an age model and sediment accumulation rates for North Pond were determined using nannofossil biostratigraphy. We found that sedimentary P was mainly present in mineral phases at both sampled locations. However, the relative abundance of different P reservoirs varied substantially as a result of different P sources, accumulation rates, redox conditions and microbial activity. This work enables a better understanding of the P cycling dynamics at each site, and the potential P sources that may be fueling deep subseafloor microbial life.

1 Introduction

1.1 Motivation and project goal

Phosphorus (P) is essential to all living cells, and the biotic and abiotic processes impacting its burial efficiency in marine sediments represent a critical component of its global biogeochemical cycle. Sources of P to marine sediments are comprised of detrital inorganic and organic material transported to the ocean by rivers or aerosols, as well as sinking biogenic material produced in the overlying water column [Paytan and McLaughlin, 2007]. Once at the sediment–water interface, recalcitrant particulate

P forms, such as detrital phosphates, are passively buried, while reactive particulate phases are subject to a number of biotic and abiotic alterations that can cause P to be released back to the water column or be retained permanently [Ruttenberg, 2014].

These alterations may also change the form in which P is ultimately buried. Such processes include microbial breakdown of organic P compounds and production of dissolved inorganic and organic P, uptake of phosphate via sorption unto mineral phases, uptake of phosphate during formation of authigenic minerals, and released of dissolved P from sediments to bottom waters [Ruttenberg, 2014]. Particulate organic

P from the photic zone is the major transport mode of reactive P to the seabed

[Delaney, 1998]. However, after burial, much of the organic P and P sorbed to iron

(Fe) oxides is released in porewater and captured by carbonate fluorapatite formation in the sediments due to organic matter respiration and changes in redox conditions, thus significantly increasing P burial efficiency [Paytan and McLaughlin, 2007].

The worldwide ridge system covers 65,000 km, making ridge flank environments a substantial area of the seafloor. They are commonly characterized by thick sediments overlying hydrologically active basaltic crust [Spinelli et al., 2004]. This environment hosts deep subseafloor microorganisms both in the sediments and upper crust, which may have profound implications on global biogeochemical cycles [Orcutt et al.,

2011; Edwards et al., 2012]. Given the prevalence of ridge flank environments, it is important to constrain the effects of parameters such a basement age, redox conditions, sediment accumulation rates, microbial activity and interactions between the upper oceanic crustal aquifer and overlying sediments on P geochemistry.

In this study, we present high-resolution solid-phase P data from two well-studied sites on the flanks of the Juan de Fuca Ridge and the Mid-Atlantic Ridge. We use the

SEDEX sequential extraction scheme [Ruttenberg, 1992], as modified by Ruttenberg et al. [2009] to gain insights into the nature of sedimentary P reservoirs, as well as the diagenetic processes influencing P at those sites. This method has been widely used in the field of marine sediment P biogeochemistry, and allows the quantification of both carbonate fluorapatite (reactive P sink) and detrital apatite (unreactive P) separately

[Ruttenberg, 1992; Delaney, 1998]. We also provide an age model for North Pond sediments derived from nannofossil biostratigraphy, as well as sediment accumulation rates for each borehole.

1.2 Field site locations

1.2.1 Mid-Atlantic Ridge Flank (IODP Expedition 336)

This study’s first field site is located at North Pond, an approximately 8 × 15 km sediment-filled basin located on the western flank of the Mid-Atlantic ridge sampled during IODP Expedition 336 [Expedition 336 Scientists, 2012]. Such sediment ponds surrounded by high relief topography are common to the western flank of the Mid-

Atlantic Ridge, and thought to be representative of slow-spreading ridge flank environments [Becker et al., 2001]. Sediments at North Pond are predominantly nannofossil ooze with layers of coarse foraminiferal sand and occasional pebble-sized clasts of basalt, serpentinite, gabbroic rocks, and bivalve debris [Expedition 336

Scientists, 2012]. Oceanic crust at this location is estimated to be 7.43 − 8.07 Ma

[Becker et al., 2001]. Oxygen penetrates deep in the sediment column, and has a “C” shaped depth profile due to oxygen fluxes from the underlying basalt crust, with the middle portion of the cores being suboxic to anoxic [Orcutt et al., 2013]. Low seafloor heat flow measurements suggest that seawater recharge into the igneous basement is occurring in the southeastern part of the basin, with fluids traveling northwestward [Langseth et al., 1992; Schmidt-Schierhorn et al., 2012]. The northwestern edge of the basin shows localized high seafloor heat flow measurements, suggestive of discharge of seawater warmed by heat flux from the lithosphere as fluids travel northwestward from the southeastern edge of the basin

[Langseth et al., 1992; Schmidt-Schierhorn et al., 2012].

1.2.2 Juan de Fuca Ridge Flank (IODP Expedition 327)

This study’s second field site is located approximately 100 km east of the Juan de

Fuca Ridge and consists of young oceanic crust (3.5–3.6 Ma) covered by thick sediments (25–600 m thick) punctuated by seamounts [Underwood et al., 2005;

Fisher et al., 2011]. This is due to high sedimentation rates and the trapping of turbidites flowing from east to west. Site U1363 was drilled during IODP Expedition

327 and is located on the edge of the Grizzly Bare seamount. Heat flow, seismic, geochemical and modeling studies have indicated that Grizzly Bare is a site of seawater recharge into the basaltic crust, and suggested a northeastward fluid flow direction towards Baby Bare outcrop, where warm, altered seawater is discharged

[Fisher et al., 2003; Hutnak et al., 2006; Wheat et al., 2013; Winslow et al., 2016].

Sediments at Hole U1363B are composed of hemipelagic mud (clayey silt to silty clay), thin-bedded turbidites (sand-silt-clay) and thickly bedded medium sand turbidite (lithostratigraphic unit 1), as well as beds of silts and sandy silt intercalated with hemipelagic mud deposits (lithostratigraphic unit 2) [Expedition 327 Scientists,

2011]. Sediment retrieved from boreholes U1363C/D consist of beds of silts and sandy silt intercalated with hemipelagic mud deposits only. Unlike the North Atlantic field site, sediments around Grizzly Bare are anoxic throughout most of the sediment column, as evidenced by porewater depth profiles of sulfate and alkalinity [Wheat et al., 2013].

2 Materials and Methods

2.1 Sample collection and storage

North Pond sediment cores were collected during IODP Expedition 336, and core recovery procedures have been described in detail [Expedition 336 scientists, 2012].

The sediment samples used for this work were retrieved from boreholes U1382B,

U1383D, U1383E, and U1384A using the advanced piston corer (Fig. 2–1a, Table 2–

1). Site U1382 is located in the southeastern part of North Pond, where sediment thickness reaches approximately 90 m. Site U1383 is located on the northeastern part of North Pond, near the edge of the basin where sediments are ~43 m thick. Site 1384 is located on the northwestern side of the basin, where sediment thickness reaches

~95 m (Table 2–1). The sediment samples used for this study were squeezed-cakes stored at –80oC after porewater collection immediately following core retrieval.

Juan de Fuca ridge flank sediment cores were collected during IODP Expedition 327, and detailed information about core recovery can be found in Fisher et al. [2011].

Sediment samples used in this study were retrieved from boreholes U1363B, C and D on the edge of Grizzly Bare outcrop using the advanced piston corer and extended core barrel coring system (Fig. 2–1b, Table 2–1). The sediment samples used for this study were squeezed-cakes stored at –80oC after porewater collection within a nitrogen-filled glove-bag at room temperature immediately following core retrieval

[Fisher et al., 2011].

2.2 Sediment sequential extraction

Solid-phase P concentrations were determined following the SEDEX sequential extraction scheme, which separates sedimentary P into 5 operationally defined categories: (1) loosely-sorbed P; (2) ferric Fe-bound P; (3) authigenic carbonate fluorapatite, biogenic apatite and calcium carbonate-associated P (CaCO3-bound P);

(4) detrital apatite and (5) organic P [Ruttenberg, 1992; Ruttenberg et al., 2009]. The appeal of the SEDEX method is that it has been successfully used in numerous studies in the field of P biogeochemistry [Ruttenberg, 2014]. Furthermore, it is the only procedure that allows for the separation of carbonate fluorapatite from the detrital apatite pool [Ruttenberg, 1992]. This is critical, since authigenic apatite represents a sink for reactive P, while detrital apatite does not [Delaney, 1998]. The separation of sedimentary P pools is based on the reactivity of each given pool to a particular extractant solution. Briefly, we lyophilized, ground minimally with an agate mortar and pestle, and sieved frozen squeezed sediment cakes to < 125 µm. We then extracted each sample (0.08 g) in 1 M magnesium chloride (MgCl2; pH 8.0) to solubilize phosphate loosely bound to mineral phases. This was followed by a reductive step (sodium citrate, dithionite and bicarbonate buffer; pH 7.6) during which sodium dithionite reduces Fe oxyhydroxides, and citrate complexes with Fe, thus releasing P associated with Fe oxides. We then washed the sediment sample residues in 1 M MgCl2 to remove any residual oxide-bound P. Next, we extracted authigenic/biogenic apatite and P bound to calcium carbonate in 1 M sodium acetate

(pH 4.0), and washed the sediment sample residues twice in 1 M MgCl2. We extracted detrital apatite in 1 M hydrochloric acid (HCl), then organic P in 1 M HCl following ashing of sample residues at 550oC [Ruttenberg et al., 2009].

2.3 Supernatant P analyses

We measured total P concentrations in sediment extracts using inductively coupled plasma optical emission spectroscopy (ICP-OES) [Perkin-Elmer Optima 4300 DV

Inductively Coupled Plasma Optical Emission Spectrometer operated by the

University of California, Santa Cruz]. Standards were prepared with the same solutions as those used for the extraction procedure in order to minimize matrix effects on P measurements, and ranged from 0 – 10 mg P L-1. The detection limit for

P on this instrument for both wavelengths is 11 µg L-1. Low P blanks associated with polycarbonate filters in the manifold reaction vessels were determined for each extraction step and sample P measurements were corrected accordingly. We used a repeat extraction of an internal sediment standard to determine the precision for each

SEDEX extraction step with analyses by inductively coupled plasma optical emission spectroscopy and assessed them to be 28%, 9%, 11%, 17% and 15% for steps 1 through 5 respectively (Table 2–2). The higher error associated with loosely-sorbed P measurements is due to the low P concentrations in those sediment extracts following sample dilution, which introduces uncertainty when measuring P concentrations on the ICP-OES.

2.4 Nannofossil biostratigraphy

A total of 121 samples were analyzed for calcareous nannofossil biostratigraphy from

IODP Expedition 336 Holes U1382B, U1383D, U1383E, and U1384A. Smear slides were made from the sediment using the methods described in Watkins and Bergen

[2003]. The slides were analyzed under a light microscope at ×1000 magnification, and fossil preservation was assessed and described as either “good,” “moderate,” or

“poor”. The Neogene zonation of Martini [1971] was used with modifications from

Expedition 320/321 Scientists [2010] (Supplementary Table S2–1). The fossil datum ages are from the Geologic Time Scale 2012 [Gradstein et al., 2012]. Average sedimentation rates were calculated by dividing the age of the oldest datum by the average depth that the datum occurs.

2.5 Phosphorus accumulation rates

We determined P accumulation rates through time by multiplying total solid-phase P by dry bulk density and the sedimentation rate derived from nannofossil biostratigraphy [Filippelli, 1997] (Supplementary Tables S2–2, S2–3). For Holes

U1363B, U1363C and U1363D, we used previously published sediment accumulation rates for hemipelagic mud and sandy turbidites [Su et al., 2000;

Underwood et al., 2005]. Dry bulk density measurements were conducted by shipboard scientists during IODP Expeditions 327 and 336 [Fisher et al., 2011;

Expedition 336 scientists, 2012]. Total solid-phase P was determined by ashing 0.3 g of sediment sample at 550oC during 2 h, then extracting samples in 30 mL of 0.5 M

sulfuric acid over 16 h [Cade-Menun and Lavkulich, 1997]. Phosphorus content in the acid extracts was determined as described above in the section “Supernatant P analyses” and reported in Supplementary Tables S2–2 and S2–3.

2.6 Statistical analyses

Statistical analyses were performed using SPSS (IBM SPSS Statistics, version 23.0).

We used a Kruskal-Wallis H test to assess potential differences in the different sedimentary P pool concentrations across the four boreholes at North Pond: U1382B

(n=42), U1383D (n=16), U1383E (n=20) and U1384A (n=28). All distributions, except those of loosely-sorbed P, were similar, as assessed boxplot inspection.

Pairwise comparisons were performed using the procedure from Dunn [1964] with a

Bonferroni correction for multiple comparisons. We also used a Kruskal-Wallis H test to determine if there were any variations in reactive P content across the three boreholes on the edge of the Grizzly Bare outcrop: U1363B (n=14), U1363C (n=4) and U1363D (n=6). Reactive P is defined at the sum of loosely-sorbed P, Fe-bound P, authigenic/biogenic apatite, CaCO3-bound P and organic P. Distributions of reactive

P content were also similar for all three boreholes. We used a one-way ANOVA to assess differences in detrital apatite content at the three boreholes at Grizzly Bare outcrop. There were no outliers, as assessed by boxplot inspection, data was normally distributed for each borehole, as assessed by the Shapiro-Wilk test (p > 0.05) and there was homogeneity of variances, as determined using Levene’s test of homogeneity of variances (p = 0.459). Data from the one-way ANOVA is presented

as mean ± standard deviation. Lastly, we used a Spearman’s rank-order correlation to assess the relationship between porewater molybdate reactive P (MRP, which includes primarily free orthophosphate), Fe, and sulfate at the Grizzly Bare site. The porewater analyses were conducted by shipboard scientists [Fisher et al., 2011].

Preliminary analysis showed the relationships to be monotonic, as determined by visual inspection of a scatterplot (results not shown).

3 Results

3.1 North Pond sedimentary P geochemistry

Sedimentary P at North Pond is mainly present in mineral phases, with the most abundant P pools being (1) P adsorbed onto Fe oxyhydroxides and (2) P present in authigenic, biogenic apatite and CaCO3-bound P (Fig. 2–2) [Defforey and Paytan,

2015]. At Hole U1382B, loosely-sorbed P represents 0–20% of total sedimentary P,

Fe-bound P 12–46%, authigenic/biogenic and CaCO3–P 6–74%, detrital P 1–24% and organic P 0–39%. At Hole U1383D, loosely-sorbed P represents 0–4% of total sedimentary P, Fe-bound P 32–45%, authigenic/biogenic and CaCO3–P 48–61%, detrital P 0–5% and organic P 0–6%. At Hole U1383E, loosely-sorbed P represents

0–9% of total sedimentary P, Fe-bound P 10–43%, authigenic/biogenic and CaCO3–P

42–77%, detrital P 2–11% and organic P 0–9%. At Hole U1384A, loosely-sorbed P represents 0–5% of total sedimentary P, Fe-bound P 26–53%, authigenic/biogenic and

CaCO3–P 35–69%, detrital P 2–22% and organic P 0–17%. Organic P is present in small amounts throughout the sediment core, even near the sediment-basement

interface (Fig. 2–2). Total sedimentary P shows little variation with depth in the sediment column, with the exception of sediment layers near the sediment-basement interface in the two long cores. Solid-phase P content is greatest at 92 mbsf at Hole

U1384A and at 89 mbsf at Hole U1382B (Fig. 2–2).

Phosphorus accumulation rates reached up to 50 µmol P cm-2 kyr-1 between 6–7.4 Ma, then decreased to 10–20 µmol P cm-2 kyr-1 over the past 6 Ma (Fig. 2–3). The higher P accumulation rates correspond to the sediments samples with the highest P content near the sediment-basement interface at boreholes U1382B and U1384A.

We assessed differences in the abundance of sedimentary P pools across the four boreholes using a Kruskal-Wallis H test, and found that median authigenic, biogenic

2 apatite, CaCO3-bound P (χ (3)= 5.070, p= 0.167) and organic P concentrations

(χ2(3)= 3.849, p= 0.278) were not significantly different between boreholes.

However, the differences in mean ranks of loosely-sorbed P concentrations were statistically significant between boreholes, χ2(3)= 60.498, p< 0.0005. Subsequent pairwise comparisons revealed statistically significant differences in loosely-sorbed P content between U1382B (mean rank: 80.07) and U1383D (23.31) (p< 0.0005),

U1382B and U1384A (32.05) (p< 0.0005), U1382B and U1383E (51.88) (p= 0.004), and between U1383D and U1383E (p=0.034), but not between U1383D and U1384A

(p= 1.000) or U1384A and U1383E (p= 0.166). We also found that median Fe-bound

P (χ2(3)= 24.800, p< 0.0005) and detrital apatite content (χ2(3)= 14.253, p=0.003)

were statistically significantly different across all four boreholes. More specifically, pairwise comparisons showed statistically significant differences in median Fe-bound

P between U1383E (median: 2.57) and U1382B (5.31) (p= 0.008), U1383E and

U1383D (5.64), U1383E and U1384A (6.96) (p< 0.0005), but not between U1382B and U1383D, or any other borehole combination. This post hoc analysis also showed statistically significant differences in median detrital P content between U1383D

(0.36) and U1382B (0.64) (p= 0.012), U1383D and U1384A (0.70) (p= 0.005), but not between U1383D and U1383E (0.46), or any other borehole combination.

3.2 Grizzly Bare seamount sedimentary P geochemistry

Detrital, authigenic, biogenic apatite and CaCO3-bound P are the dominant sedimentary P pools at Site U1363 on the edge of the Grizzly Bare outcrop (Fig. 2–4).

At Hole U1363B, nearest the seamount, loosely-sorbed P represents 0–4% of total sedimentary P, Fe-bound P 0–14%, authigenic/biogenic and CaCO3–P 14–48%, detrital P 31–77% and organic P 7–9%. In the deeper sediments of Hole U1363C, loosely-sorbed P represents 1–4% of total sedimentary P, Fe-bound P 0%, authigenic/biogenic and CaCO3–P 21–42%, detrital P 50–72% and organic P 5–9%.

At Hole U1363D, loosely-sorbed P represents 0–2% of total sedimentary P, Fe-bound

P 0%, authigenic/biogenic and CaCO3–P 12–42%, detrital P 53–81% and organic P

5–8%. Unlike North Pond, this site contains little to no P bound to Fe oxyhydroxides.

Iron-bound P is present in small amounts at Hole U1363B (0.6–3.2 µmol g-1), but decreases as a function of depth in the sediment column as Fe-oxyhydroxides become

reduced in the anoxic sediments. Also, there is no peak in solid-phase P content near the sediment-basement interface as observed at the X336 field sites. Total P content does not change with depth, and the proximity to the sediment-basement interface does not appear to affect sedimentary P geochemistry.

Results from a Kruskal-Wallis H test showed that median reactive P content decreased from U1363B (7.87), U1363C (7.64) to U1363D (7.04). However, the differences were not statistically significantly different, χ2(2)= 1.19, p = 0.550. We compared detrital apatite content using a one-way ANOVA and found that detrital P was similar for all three boreholes U1363B (mean ± standard deviation: 12.7±3.2),

U1363C (13.1±2.6) and U1363D (12.7±2.7), and the differences in detrital P content between the boreholes were not statistically significant, F(2,21)= 0.048, p = 0.954.

Shore-based porewater MRP measurements show a sharp increase in porewater phosphate content in the first 16 meters of the sediment column, followed by a steady decrease to values less than 1 µM [Expedition 327 Scientists, 2011]. This is concurrent with a peak in porewater Fe, DOC and alkalinity, and a decrease in porewater sulfate content [Expedition 327 Scientists, 2011; Lin et al., 2015]. There was a significant positive correlation between porewater MRP and Fe concentrations, rs(17)= 0.606 (p= 0.006) and a significant negative correlation between porewater

MRP and sulfate, rs(17)= –0.601, (p= 0.006). However, there was no relationship between porewater MRP and solid-phase organic P, or MRP and Fe-bound P.

Molar ratios of C/Porg (142±56, n=26) were variable with depth, but overall greater than the Refield ratio (106; Table 2–3) [Redfield, 1934]. Molar ratios of C/Preact were lower (24±14, n=26) and also variable with depth. Hole U1363C had the lowest C/P ratios for both organic and reactive P, while Hole U1363D had the highest ratios.

Phosphorus accumulation from 242 µmol cm-2 kyr-1 for hemipelagic muds, to 281–

1330 µmol cm-2 kyr-1 for sandy turbidites (Supplementary Table S2–3). Sediment accumulation rates are the main factor controlling the magnitude of P accumulation at this site.

4 Discussion

4.1 North Pond sedimentary P cycling

Sedimentary P at North Pond is predominantly present in mineral phases, with 6–74% of total P present in authigenic, biogenic and CaCO3-bound P and 12–53% bound to

Fe oxyhydroxides. Since most of the sediment column is oxic [Orcutt et al., 2013], both authigenic apatite and Fe-oxide bound P act as a major burial sinks for P and no significant sink switching occurs between those two reservoirs [Ruttenberg, 2014].

While authigenic apatite is known to be a major sink [Ruttenberg, 2014], Fe-bound P has also been shown to act as a burial sink for P [Slomp et al., 2013]. There is no significant difference in organic P content across all boreholes. This is consistent with findings from the North Pond site survey, which showed low sedimentary organic C

(0.15±0.07%) and nitrogen (0.03±0.05%) content with no discernable differences

across the sampled sites [Ziebis et al., 2012]. Organic P abundance decreases with depth, consistent with organic matter remineralization. However, at Sites U1382 and

U1384, organic P along with most sedimentary P pools except loosely sorbed P, show a localized increase in abundance near the sediment–basement interface.

Total solid-phase P concentrations measured fall within the range of concentrations measured for other open ocean regions [Filippelli, 1997] and show little variation with depth, except near the sediment-basement interface at boreholes U1382B and

U1384A, where they increase substantially. There is no indication of P diffusive fluxes from the basement, as evidenced by porewater phosphate concentrations, which range from 0.81–1.07 µM P near the sediment–basement interface [Expedition

336 scientists, 2012]. The increase in sedimentary P content near the interface at boreholes U1382B and U1384A could be due to a past increase in export production to the seafloor. This is supported by higher P accumulation rates between 6–7.4 Ma

(Fig. 2–3). A possible explanation for this increase in export production could be higher primary productivity in the overlying water column at the time this sediment layer was deposited. The decrease in P accumulation rates from 7.4 Ma to 6 Ma could be due to a gradual change in oceanographic regime. One possible explanation for this is the closing of the Panama gateway, which was initiated 17 Ma [Brierley and

Fedorov, 2016] and resulted in the modern Caribbean oceanographic regime starting

6–8 Ma [Collins et al., 1996]. Changes in benthic foraminiferal species indicate that the closing of the Panama gateway resulted in a decrease in primary productivity in

the North Atlantic [Schmittner et al., 2004]. In addition, simulations have shown a decrease in nutrient concentrations and a net decrease in primary productivity in the

North Atlantic as a result of the gradual closing of the gateway, consistent with sediment records and supporting the hypothesis that primary productivity in the North

Atlantic was strongly dependent on nutrients from the East Pacific [Schneider and

Schmittner, 2006]. One way to test this hypothesis would be to assess the biogenic barite content in the sediment layers where P maxima are observed near the sediment–basement interface. Biogenic barite is linked to particulate organic C and primary productivity, well preserved in oxic sediments and is a paleoceanographic proxy that can be used to reconstruct export production [Paytan and Griffith, 2007].

We found a statistically significant difference in detrital apatite content between

Holes U1383D and U1382B, U1383D and U1384A but not between boreholes at Site

U1383. A possible explanation for this is the location of these sites and the frequency of sediment flows they may experience. Indeed, sediment samples from Site U1383, especially U1383D samples, showed more reworking (possibly turbidites) upon visual inspection under a microscope. Site U1383 is located on the northeastern part of North Pond, near the edge of the basin. Sites U1382 and U1384 are located on the southeastern and northwestern sides of the basin respectively, further away from the basin edges. As a result of the different locations along the basin, those two sites may have experienced less sediment flows that could account for the differences in detrital apatite abundance.

4.2 Grizzly Bare seamount solid-phase P cycling

Similarly to results from North Pond, sedimentary P from Site U1363 is dominantly present in mineral phases (Fig. 2–4). While authigenic, biogenic apatite and CaCO3- bound P dominate North Pond solid-phase P, U1363 sedimentary P consists mainly of detrital apatite (31–81%) and, to a lesser extent, of authigenic, biogenic apatite and

CaCO3-bound P (12–48%; Fig. 2–4). Thus, the dominant sink of reactive P in this system is authigenic apatite. Total P varies little with depth, and Fe-bound P is present in small amounts in shallower sediments, but is absent deeper in the sediment column. This is expected given the anoxic nature of these sediments [Fisher et al.,

2011], which causes ferric iron oxides to be reduced and adsorbed P to be released in porewater.

The significant positive correlation between porewater MRP and Fe, and the significant negative correlation between MRP and sulfate suggest that the increase in porewater MRP observed in the first 16 mbsf could be related to both (1) MRP release by organic matter remineralization through sulfate reduction, and (2) release of MRP adsorbed onto Fe(III) oxyhydroxides from Fe oxide reduction (Table 2–3)

[Filippelli and Delaney, 1996; Paytan and McLaughlin, 2007; Ruttenberg, 2014]. Lin et al. [2015] reported a correlation between porewater sulfate and dissolved organic carbon (DOC), and suggested that the increase in porewater DOC in the first 16 mbsf could be related to sedimentary particulate organic matter remineralization. Sulfate reduction produces sulfide, which contributes to Fe(III) oxyhydroxide dissolution and

also likely enhances P remineralization from organic matter [Mort et al., 2010]. We did not find a correlation between POC and POP, or POC and Preact which has been observed in a number of different marine sediments [Ingall and Van Cappellen, 1990;

Anderson et al., 2001].

Molar ratios of C/P varied with depth, but on average C/Porg ratios were greater than the Redfield ratio, while C/Preact were substantially lower. Higher C/Porg ratios suggest enhanced P remineralization from organic matter relative to C or a terrestrial source of organic matter [Ruttenberg and Goñi, 1997]. However, while C/Porg ratios are valuable, C/Preact ratios are a more appropriate proxy for describing sedimentary P geochemical behavior since they capture the efficient transfer of loosely-sorbed P, iron-bound P and organic P to authigenic apatite [Anderson et al., 2001]. Molar

C/Preact ratios are substantially lower than the Redfield ratio, which indicates that preferential P release is not occurring at Site U1363. It is worth noting that higher C/P ratios were observed in sandy intervals corresponding to major turbidite events (Table

2–3). Lin et al. [2015] found that C/N ratios were slightly higher than the Redfield ratio (6.6), but significantly lower than that of terrestrial vascular plants (>20)

[Redfield, 1934; Meyers, 1994]. The higher C/P ratios in turbidite layers support the conclusion from Lin et al. [2015] that sediments at Site U1363 are likely a mixture of planktonic biomass and terrestrial debris.

Detrital phosphates are passively buried in sediments [Ruttenberg, 2014] and originate primarily from land [Jaisi and Blake, 2010]. Underwood et al. [2005] investigated the provenance and stratigraphy of sediments on the flank of the Juan de

Fuca Ridge flank, but were unable to identify a unique sand provenance. They concluded that a mixed provenance contributed sand to the northwestern part of the

Cascadia Basin from Vancouver Island, the Olympic Peninsula, the northern

Cascades and western British Columbia. Erosion of andesitic and rhyolitic rock, as well as ash deposits in those watersheds delivers detrital apatite to the continental margin. From there, the trapping of turbidites flowing from east to west by topographic relief associated with the Juan de Fuca Ridge axis and the abyssal hill bathymetry on the ridge flank would allow the accumulation of detrital apatite in sediments [Fisher et al., 2011].

5 Summary and conclusions

We present high-resolution solid-phase P data from two well-studied field sites sampled during IODP expeditions 327 and 336. Site U1363 is located on the edge of the Grizzly Bare seamount, in the Juan de Fuca Ridge flank area, and is characterized by young oceanic crust (~3.5 Ma), thick anoxic sediments and high sedimentation rates. Sedimentary P at this site is mainly present in the form of detrital apatite, likely transported by turbidity currents from east to west. Iron-bound P is rapidly reduced in these sediments, and the second most abundant solid-phase P reservoir is

authigenic/biogenic apatite and CaCO3-bound P. In contrast, the North Atlantic field site North Pond has oxic sediments of varying thickness overlying slightly older oceanic crust (7.4–8.1 Ma). Sedimentary P at this site is primarily in the form of authigenic/biogenic apatite and CaCO3-bound P, and Fe-bound P. The latter form persists at depth due to the oxygenated sediments at the top and bottom of the sediment column. Unlike Site U1363, the sites at North Pond have little detrital apatite due to their distance from continental margins. The difference in P accumulation rates between the sites is mainly due to differences in proximity to a continent and associated runoff, with marine snow being the main P source to North

Pond sediments. A past increase in export production is a possible explanation for the localized increase in P near the sediment-basement interface.

We show that sedimentary P geochemistry varies from one ridge flank system to another, and that the SEDEX sequential extraction scheme yields valuable information regarding the nature and abundance of different P reservoirs and their relative reactivity. This is of relevance to ongoing microbiology studies at both locations, as this dataset provides insights into possible P sources to the deep biosphere. We also show that organic P is present at both the Juan de Fuca Ridge and

Mid-Atlantic Ridge flank sites and, depending on the degree of sorption on mineral surfaces and redox conditions, may constitute a bioavailable P source to deep sedimentary subseafloor microorganisms.

Acknowledgments

We thank the Integrated Ocean Drilling Program, and the shipboard scientists and technical staff involved in IODP expeditions 327 and 336 who collected the sediment samples used in this study. Funding was provided by a graduate student fellowship from the Center for Dark Energy Biosphere Investigations (NSF C-DEBI) to D.D.

(award # 157598), and a research grant from C-DEBI to A.P. (award # 156246). We also thank Rob Franks, Michael Kong, Caitriona Berger and Natalie Zimdahl for their assistance with this project.

References

Anderson, L. D., M. L. Delaney, and K. L. Faul (2001), Carbon to phosphorus ratios in sediments: implications for nutrient cycling, Glob. Biogeochem. Cycles, 15(1), 65–79, doi:10.1029/2000GB001270.

Becker, K., A. Bartetzko, and E. E. Davis (2001), Leg 174B synopsis: revisiting Hole 395A for logging and long-term monitoring of off-axis hydrothermal processes in young oceanic crust, Proc. ODP, 174B, doi:10.2973/ odp.proc.sr.174B.130.2001.

Brierley, C. M., and A. V. Fedorov (2016), Comparing the impacts of Miocene– Pliocene changes in inter-ocean gateways on climate: Central American Seaway, Bering Strait, and Indonesia, Earth Planet. Sci. Lett., 444, 116–130, doi:10.1016/j.epsl.2016.03.010.

Cade-Menun, B. J., and L. M. Lavkulich (1997), A comparison of methods to determine total, organic, and available phosphorus in forest soils, Commun. Soil Sci. Plant Anal., 28(9-10), 651–663, doi:10.1080/00103629709369818.

Collins, L. S., A. G. Coates, W. A. Berggren, M.-P. Aubry, and J. Zhang (1996), The late Miocene Panama isthmian strait, Geology, 24(8), 687–690, doi:10.1130/0091-7613(1996)024<0687:TLMPIS>2.3.CO;2.

Defforey, D., and A. Paytan (2015), Data report: characteristics of sedimentary phosphorus at North Pond, IODP Expedition 336, Proc. Integr. Ocean Drill. Program, 336, doi:10.2204/iodp.proc.336.205.2015.

Delaney, M. L. (1998), Phosphorus accumulation in marine sediments and the oceanic phosphorus cycle, Glob. Biogeochem. Cycles, 12(4), 563–572, doi:10.1029/98GB02263.

Dunn, O. J. (1964), Multiple Comparisons Using Rank Sums, Technometrics, 6(3), 241–252, doi:10.1080/00401706.1964.10490181.

Edwards, K. J., K. Becker, and F. Colwell (2012), The deep, dark energy biosphere: intraterrestrial life on earth, Annu. Rev. Earth Planet. Sci., 40(1), 551–568, doi:10.1146/annurev-earth-042711-105500.

Expedition 320/321 Scientists (2010), Methods, IODP Proc., 320/321, doi:10.2204/iodp.proc.320321.102.2010.

Expedition 327 Scientists (2011), Site U1363, IODP Proc., 327, doi:10.2204/iodp.proc.327.106.2011.

Expedition 336 Scientists (2012), Initiation of long-term coupled microbiological, geochemical, and hydrological experimentation within the seafloor at North Pond, western flank of the Mid-Atlantic Ridge., IODP Proc., 336, doi:10.2204/iodp.pr.336.2012.

Expedition 336 scientists (2012), Sediment and basement contact coring, Proc. Integr. Ocean Drill. Program, 336, doi:10.2204/iodp.proc.336.106.2012.

Filippelli, G. M. (1997), Controls on phosphorus concentration and accumulation in oceanic sediments, Mar. Geol., 139, 231–240, doi:10.1016/S0025- 3227(96)00113-2.

Filippelli, G. M., and M. L. Delaney (1996), Phosphorus geochemistry of equatorial Pacific sediments, Geochim. Cosmochim. Acta, 60(9), 1479–1495, doi:10.1016/0016-7037(96)00042-7.

Fisher, A. T. et al. (2003), Hydrothermal recharge and discharge across 50 km guided by seamounts on a young ridge flank, Nature, 421(6923), 618–621, doi:10.1038/nature01352.

Fisher, A. T., T. Tsuji, K. Petronotis, and Expedition 327 Scientists (2011), IODP Proc., 327, Tokyo (Integrated Ocean Drilling Program Management International, Inc.), doi:10.2204/iodp.proc.327.2011.

Gradstein, F. M., J. G. Ogg, M. D. Schmitz, and G. M. Ogg (2012), The Geologic Time Scale 2012, Elsevier, Amsterdam.

Hutnak, M., A. T. Fisher, L. Zühlsdorff, V. Spiess, P. H. Stauffer, and C. W. Gable (2006), Hydrothermal recharge and discharge guided by basement outcrops on 0.7–3.6 Ma seafloor east of the Juan de Fuca Ridge: Observations and numerical models, Geochem. Geophys. Geosystems, 7(7), Q07O02, doi:10.1029/2006GC001242.

Ingall, E. D., and P. Van Cappellen (1990), Relation between sedimentation rate and burial of organic phosphorus and organic carbon in marine sediments, Geochim. Cosmochim. Acta, 54(2), 373–386, doi:10.1016/0016- 7037(90)90326-G.

Jaisi, D. P., and R. E. Blake (2010), Tracing sources and cycling of phosphorus in Peru Margin sediments using oxygen isotopes in authigenic and detrital phosphates, Geochim. Cosmochim. Acta, 74(11), 3199–3212, doi:10.1016/j.gca.2010.02.030.

Langseth, M. G., K. Becker, R. P. Vonherzen, and P. Schultheiss (1992), Heat and fluid flux through sediment on the western flank of the Mid-Atlantic Ridge: a hydrogeological study of North Pond, Geophys. Res. Lett., 19(5), 517–520, doi:10.1029/92GL00079.

Lin, H.-T., C.-C. Hsieh, J. P. Cowen, and M. S. Rappé (2015), Data report: dissolved and particulate organic carbon in the deep sediments of IODP Site U1363 near Grizzly Bare seamount, Proc. Integr. Ocean Drill. Program, 327, doi:10.2204/iodp.proc.327.202.2015.

Martini, E. (1971), Standard Tertiary and Quaternary calcareous nannoplankton zonation, in Proceedings of the second Planktonic Conference, edited by A. Farinacci, pp. 739–785, Edizioni Tecnoscienza Roma.

Meyers, P. A. (1994), Preservation of elemental and isotopic source identification of sedimentary organic matter, Chem. Geol., 114(3), 289–302, doi:10.1016/0009-2541(94)90059-0.

Mort, H. P., C. P. Slomp, B. G. Gustafsson, and T. J. Andersen (2010), Phosphorus recycling and burial in Baltic Sea sediments with contrasting redox conditions, Geochim. Cosmochim. Acta, 74(4), 1350–1362, doi:10.1016/j.gca.2009.11.016.

Orcutt, B. N., J. B. Sylvan, N. J. Knab, and K. J. Edwards (2011), Microbial Ecology of the Dark Ocean above, at, and below the Seafloor, Microbiol. Mol. Biol. Rev., 75(2), 361–422, doi:10.1128/MMBR.00039-10.

Orcutt, B. N., C. G. Wheat, O. Rouxel, S. Hulme, K. J. Edwards, and W. Bach (2013), Oxygen consumption rates in subseafloor basaltic crust derived from a reaction transport model, Nat. Commun., 4, doi:10.1038/ncomms3539.

Paytan, A., and E. M. Griffith (2007), Marine barite: Recorder of variations in ocean export productivity, Deep Sea Res. Part II Top. Stud. Oceanogr., 54(5–7), 687–705, doi:10.1016/j.dsr2.2007.01.007.

Paytan, A., and K. McLaughlin (2007), The Oceanic Phosphorus Cycle, Chem. Rev., 107(2), 563–576, doi:10.1021/cr0503613.

Redfield, A. C. (1934), On the proportions of organic derivatives in sea water and their relation to the composition of plakton, in James Johnson Memorial Volume, edited by R. J. Daiel, pp. 177–192, University Press Liverpool, Liverpool.

Ruttenberg, K. C. (1992), Development of a sequential extraction method for different forms of phosphorus in marine sediments, Limnol. Oceanogr., 37(7), 1460–1482, doi:10.4319/lo.1992.37.7.1460.

Ruttenberg, K. C. (2014), The Global Phosphorus Cycle, in Treatise on Geochemistry, vol. 10, edited by H. Holland and K. Turekian, pp. 499–558, Elsevier.

Ruttenberg, K. C., and M. A. Goñi (1997), Phosphorus Cycling and Sedimentation in Marine and Freshwater SystemsPhosphorus distribution, C:N:P ratios, and δ13Coc in arctic, temperate, and tropical coastal sediments: tools for characterizing bulk sedimentary organic matter, Mar. Geol., 139(1), 123–145, doi:10.1016/S0025-3227(96)00107-7.

Ryan, W. B. F. et al. (2009), Global Multi-Resolution Topography synthesis, Geochem. Geophys. Geosystems, 10(3), Q03014, doi:10.1029/2008GC002332.

Schmidt-Schierhorn, F., N. Kaul, S. Stephan, and H. Villinger (2012), Geophysical site survey results from North Pond (Mid-Atlantic Ridge), IODP Proc., 336, doi:10.2204/iodp.proc.336.107.2012.

Schmittner, A. et al. (2004), Global impact of the Panamanian Seaway, Eos Trans. Am. Geophys. Union, 85(49), 526–527, doi:10.1029/2004EO490010.

Schneider, B., and A. Schmittner (2006), Simulating the impact of the Panamanian seaway closure on ocean circulation, marine productivity and nutrient cycling, Earth Planet. Sci. Lett., 246(3-4), 367–380, doi:10.1016/j.epsl.2006.04.028.

Slomp, C. P., H. P. Mort, T. Jilbert, D. C. Reed, B. G. Gustafsson, and M. Wolthers (2013), Coupled Dynamics of Iron and Phosphorus in Sediments of an Oligotrophic Coastal Basin and the Impact of Anaerobic Oxidation of Methane, PLOS ONE, 8(4), e62386, doi:10.1371/journal.pone.0062386.

Spinelli, G. A., E. R. Giambalvo, and A. T. Fisher (2004), Sediment permeability, distribution, and influence on fluxes in oceanic basement, in Hydrogeology of the Oceanic Lithosphere, edited by E. E. Davis and H. Elderfield, pp. 151– 188, Cambridge University Press, Cambridge, UK.

Su, X., K. H. Baumann, and J. Thiede (2000), Calcareous nannofossils from leg 168: biochronology and diagenesis, in Proceedings of the Ocean Drilling Program, Scientific Results, vol. 168, pp. 39–49.

Underwood, M. B., K. D. Hoke, A. T. Fisher, E. E. Davis, E. Giambalvo, L. Zuhlsdorff, and G. A. Spinelli (2005), Provenance, Stratigraphic Architecture, and Hydrogeologic Influence of Turbidites on the Mid-Ocean Ridge Flank of Northwestern Cascadia Basin, Pacific Ocean, J. Sediment. Res., 75(1), 149– 164, doi:10.2110/jsr.2005.012.

Watkins, D. K., and J. A. Bergen (2003), Late Albian adaptive radiation in the calcareous nannofossil genus Eiffellithus, Micropaleontology, 49(3), 231–251, doi:10.2113/49.3.231.

Wheat, C. G., S. M. Hulme, A. T. Fisher, B. N. Orcutt, and K. Becker (2013), Seawater recharge into oceanic crust: IODP exp 327 site U1363 Grizzly Bare outcrop: seawater recharge into basaltic crust, Geochem. Geophys. Geosystems, 14(6), 1957–1972, doi:10.1002/ggge.20131.

Winslow, D. M., A. T. Fisher, P. H. Stauffer, C. W. Gable, and G. A. Zyvoloski (2016), Three-dimensional modeling of outcrop-to-outcrop hydrothermal circulation on the eastern flank of the Juan de Fuca Ridge: Three-Dimensional Ridge-Flank Hydrothermal Modeling, J. Geophys. Res. Solid Earth, 121(3), 1365–1382, doi:10.1002/2015JB012606.

Ziebis, W., J. McManus, T. Ferdelman, F. Schmidt-Schierhorn, W. Bach, J. Muratli, K. J. Edwards, and H. Villinger (2012), Interstitial fluid chemistry of sediments underlying the North Atlantic gyre and the influence of subsurface fluid flow, Earth Planet. Sci. Lett., 323-324, 79–91, doi:10.1016/j.epsl.2012.01.018.

3100 3400 3800 4100 4500 2200 2400 2600 2800 Depth (mbsl) 3600 Depth (mbsl)

3800 18’ 52’ o o 4200 U1363C, D

U1363B U1384A

48’ 22 4000 o U1383D,E 4400 2200

2300 U1382B 16’ N 16’ 47

o 2400 2600 47 44’ N 44’ 22 o 22

0 1 2 km (a)A 0 1 2 km (b)

46o 10’ W 46o 5’ 46o 0’ 128o 06’ W 128o 02’ Fig. 2-1. Location of the Mid-Atlantic Ridge flank field site (North Pond, panel a) and the Juan de Fuca Ridge flank field site (Grizzly Bare seamount, panel b). The color scales indicate water depth for each site in meters below sea level (mbsl). Contour line intervals are 100 m for North Pond and 50 m for Grizzly Bare seamount. Location of coring sites are indicated with white dotes. Map created using the default Global Multi-Resolution Topography Synthesis basemap [Ryan et al., 2009] in GeoMapApp version 3.6.0 (http://www.geomapapp.org).

P concentration (μmol g-1) 02405100510024012 0

40 U1382B U1383D 50 U1383E U1384A Depth (mbsf) Depth 60

90 (a) (b) (c) (d) (e) 100 051002040020400 102001020 P concentration (μmol g-1) Fig. 2-2. Solid-phase P concentrations at North Pond determined using the SEDEX sequential extraction scheme, with each panel representing a different sedimentary P pool: (a) loosely-sorbed P, (b) Fe-bound P, (c) authigenic/biogenic apatite + CaCO3- bound P, (d) detrital apatite and (e) organic P. The horizontal axis at the top of the figure corresponds to P concentration scale for the first 85 mbsf (meters below seafloor). The bottom horizontal axis (in blue) corresponds to the P concentration scale for depths greater than 85 mbsf. Basement depths are 90 mbsf (U1382B), 43.3 mbsf (U1383D), 43.2 mbsf (U1383E) and 94.7 mbsf (U1384A).

P acc. rate (μmol cm-2 kyr-1) 01020304050 0 U1382B U1383D 1 U1383E U1384A 2

4 Age (Ma) 5

8 Fig. 2–3. Phosphorus accumulation rates for all four boreholes at North Pond as a function of sediment age, determined using an age-model derived from nannofossil biostratigraphy, dry bulk density measurements [Expedition 336 scientists, 2012] and total solid-phase P content.

P concentration (μmol g-1) 0 0.5 1 0 2 4 0 10 200 10 20 0 2 4 0

100

150 Depth (mbsf)

200

(a) (b) (c) (d) (e) 250 1363B 1363C 1363D

Fig. 2–4. Solid-phase P concentrations at Grizzly Bare determined using the SEDEX sequential extraction scheme, with each panel representing a different sedimentary P pool: (a) loosely-sorbed P, (b) Fe-bound P, (c) authigenic/biogenic apatite + CaCO3- bound P, (d) detrital apatite and (e) organic P. The gray boxes represent sections of the cores that were not retrieved during drilling. Basement depths are 57 mbsf (U1363B) and 231 mbsf (U1363D). Basement depth is not available for Hole U1363C.

Table 2-1. Hole locations and specifications

Seafloor Basement Boreholes Latitude Longitude deptha (mbsl) deptha (mbsf) b IODP 336 Hole 1382B 22°45.3528′N 46°04.8748 ′W 4482.5 90 Hole 1383D 22°48.1316′N 46°03.1628′W 4413.8 43 Hole 1383E 22°48.1283′N 46°03.1582′W 4413.7 43 Hole 1384A 22°48.7086′N 46°05.3464′W 4464.4 95

IODP 327c

Hole 1363B 47°17.3518′N 128°2.1060′W 2678.6 57 Hole 1363C 47°17.5759′N 128°1.7641′W 2677.6 NA Hole 1363D 47°17.5724′N 128°1.7599′W 2677.6 231 aAbbreviations: mbsl, meters below sea level; mbsf, meters below seafloor b[Expedition 336 scientists, 2012] c[Expedition 327 Scientists, 2011]

Table 2-2. Reproducibility for each SEDEX extraction step, determined by replicate analysis of an internal sediment standard (n= 16).

SEDEX step P fractiona Mean P (µmol g-1) SDa Relative errorb (%)

I Pex 0.51 0.15 28

II PFe 8.75 0.76 9

III PCFA 38.59 4.38 11

IV PDetr 31.33 5.31 17

V POrg 5.92 0.91 15 c TIP (I-IV) PInorg 79.18 3.58 5 d TP (I-V) PTot 85.10 3.48 4 a Abbreviations: Pex, loosely-sorbed/exchangeable P; PFe, P bound to iron oxyhydroxides; PCFA, authigenic carbonate fluorapatite, biogenic apatite and calcium carbonate-associated P; PDetr, detrital apatite; POrg, organic P; PInorg, inorganic P; PTot, total P. bRelative error: (SD/Mean) × 100 cTotal inorganic P (TIP) is the sum of P content from steps I-IV dTotal P (TP) is the sum of P content from steps I-V

Table 2-1. Pore water concentrations and C to P molar ratios for Site U1363 sediments.

Average Porewater Porewater Porewater Core, section Porewater Sediment Sediment Sediment depth sulfate DOC MRP interval (m) Fe (µM)a C/Nb C/P C/P (mbsf) (mM)a (mM)b (µM)a org react

327-U1363B- 1H-1, 1.4–1.5 1.5 27.8 – 13.2 8.33 10.2 59.8 13.3 2H-3, 6.9–7 7.0 23.6 0.64 – – 11.7 163.9 22.8 2H-6, 11.4–11.5 11.5 22 – 65.6 64.3 11.1 141.2 16.6 3H-3, 16.4–16.5 16.5 21.5 – 66 37.4 11.4 85.8 20.3 3H-6, 20.9–21 21.0 20.9 – 54 31.7 11.0 180.7 24.0 4H-3, 25.9–26* 26.0 21.2 – 18.1 16 10.4 141.0 17.3 4H-5, 28.9–29* 29.0 21.9 – 12.8 43 10.4 179.1 35.5 5H-2, 33.9–34 34.0 24 – 14.2 18.1 12.3 103.6 17.7 7X-1, 43.8–44 43.9 26.1 – – 20.6 15.7 82.3 24.0 7X-2, 45.3–45.5 45.4 25.8 0.35 1.99 19.6 10.2 112.8 25.7 8X-1, 48.8–49 48.9 26.4 – 4.29 16.2 152.5 27.7

8X-2, 50.3–50.5 50.4 26.6 0.35 3.41 5.48 10.7 175.9 20.4 8X-3, 51.8–52* 51.9 27.2 – – 7.62 11.6 85.2 22.6 8X-5, 53.2–53.4 53.3 27.5 – 0.87 0.29 9.3 164.0 18.8

327-U1363C- 3X-1, 173.9– 174.2 25.9 – 0.63 3.54 11.2 50.5 8.5 174.1 3X-CC, 174.1– 174.0 25.8 0.26 – 8.8 10.2 110.6 12.0 174.3 4X-1, 184.3– 184.4 26.3 0.21 1.93 24.6 10.3 164.2 27.7 184.5 4X-2, 185.8–186 185.9 26.9 0.21 1.69 18.1 12.0 127.6 15.1 5X-1, 193.1–193.3* 193.2 27.2 0.18 7.09 16.1 82.5 17.3

327-U1363D- 2X-2, 199.7– 199.8 26.8 – 0.69 20.5 11.3 158.4 17.2 199.9 3X-2, 205–205.1 205.0 26.5 0.12 0.63 9.83 12.6 233.3 86.5 3X-3, 206.5– 206.7 26.2 0.23 0.51 4.43 19.3 182.1 31.8 206.9 3X-4, 207.1– 207.3 26.8 – 0.48 6.41 16.1 117.4 18.6 207.5 4X-1, 213.1– 213.3 27.1 0.17 – – 10.8 308.5 29.3 213.5 4X-2, 214.6–215 214.8 27.7 0.16 0.69 16.6 14.1 193.5 32.3 5X-1, 222.5– 222.7 27.3 0.10 0.40 8.7 17.4 135.9 21.8 222.9 aData from Expedition 327 scientists [2011] bData from Lin et al. [2015] *Turbidite layers

Table S2-1. North Pond nannofossil biostratigraphy

Depth Nanno Sample Preservationa Age Youngest date (Ma) Oldest Date (Ma) (mbsf) Zone

Hole U1382B U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 1.5 G NN19 1H-2 IWA Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 3.1 G NN19 1H-3 IWA Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 4.6 G NN19 1H-4 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 5.6 G NN19 1H-5 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 7.37 G NN19 2H-2 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 8.87 M NN19 2H-3 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 10.17 M NN19 2H-4 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Pseudoemiliania T Helicosphaera sellii 11.27 G NN19 2H-5 Pleistocene lacunosa (0.44) (1.26) U1382B- late Pliocene/early T Helicosphaera T Calcidiscus macintyrei 13.37 M NN19 2H-6 Pleistocene sellii (1.26) (1.60) U1382B- late Pliocene/early T Helicosphaera T Calcidiscus macintyrei 14.87 M NN19 2H-7 Pleistocene sellii (1.26) (1.60) U1382B- late Pliocene/early T Helicosphaera T Calcidiscus macintyrei 18.7 M-G NN19 3H-3 Pleistocene sellii (1.26) (1.60) U1382B- late Pliocene/early T Helicosphaera T Calcidiscus macintyrei 19.5 M-G NN19 3H-4 Pleistocene sellii (1.26) (1.60) U1382B- late Pliocene/early T Calcidiscus T Discoaster brouweri 22.4 M NN19 3H-5 Pleistocene macintyrei (1.60) (1.93) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 26.15 M early Pliocene pseudoumbilicus 4H-2 IWB 15 (5.05) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 29.15 P early Pliocene pseudoumbilicus 4H-4 15 (5.05) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 32.25 M early Pliocene pseudoumbilicus 4H-6 15 (5.05) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 46.64 M early Pliocene pseudoumbilicus 6H-3 15 (5.05) (3.70) U1382B- NN13/ T Discoaster B Ceratolithus rugosus 50.78 M middle Pliocene 6H-7 15-16 tamalis (2.80) (5.05) U1382B- T Discoaster T Reticulofenestra 56.61 M NN16 middle Pliocene 7H-3 tamalis (2.80) pseudoumbilicus (3.70) U1382B- T Discoaster T Reticulofenestra 58.11 M NN16 middle Pliocene 7H-4 IWB tamalis (2.80) pseudoumbilicus (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 59.61 M early Pliocene pseudoumbilicus 7H-5 15 (5.12) (3.70) U1382B- T Discoaster T Reticulofenestra 61.11 M NN16 middle Pliocene 7H-6 tamalis (2.80) pseudoumbilicus (3.70) U1382B- T Discoaster T Reticulofenestra 62.71 M-G NN16 middle Pliocene 7H-7 tamalis (2.80) pseudoumbilicus (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 64.45 P-M early Pliocene pseudoumbilicus 8H-2 15 (5.12) (3.70)

T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 65.55 P-M early Pliocene pseudoumbilicus 8H-3 IWB 15 (5.12) (3.70) U1382B- NN13/ T Discoaster B Ceratolithus rugosus 67.45 P middle Pliocene 8H-4 IWB 15-16 tamalis (2.80) (5.12) U1382B- T Discoaster T Reticulofenestra 68.35 M NN16 middle Pliocene 8H-5 tamalis (2.80) pseudoumbilicus (3.70) U1382B- T Discoaster T Reticulofenestra 70.45 M NN16 middle Pliocene 8H-6 tamalis (2.80) pseudoumbilicus (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 71.95 M early Pliocene pseudoumbilicus 8H-7 15 (5.12) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 73.98 M early Pliocene pseudoumbilicus 9H-2 15 (5.12) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 75.48 P-M early Pliocene pseudoumbilicus 9H-3 15 (5.12) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 76.98 P-M early Pliocene pseudoumbilicus 9H-4 IWA 15 (5.12) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 78.68 P early Pliocene pseudoumbilicus 9H-5 IWB 15 (5.12) (3.70) T Reticulofenestra U1382B- NN13/ B Ceratolithus rugosus 79.98 P early Pliocene pseudoumbilicus 9H-6 IWA 15 (5.12) (3.70) U1382B- T Nicklithus B Nicklithus amplificus 81.68 P-M NN11 late Miocene 9H-7-IWB amplificus (5.94) (6.91) T Discoaster U1382B- B Amaurolithus spp. 82.91 P NN11 late Miocene quinqueramus 10H-2 (7.42) (5.59) U1382B- T Nicklithus B Nicklithus amplificus 84.41 P NN11 late Miocene 10H-3 amplificus (5.94) (6.91) U1382B- T Nicklithus B Nicklithus amplificus 10H-4 86.11 P NN11 late Miocene amplificus (5.94) (6.91) IWB U1382B- B Nicklithus B Amaurolithus spp. 87.41 P NN11 late Miocene 10H-5 amplificus (6.91) (7.42) U1382B- B Nicklithus B Amaurolithus spp. 10H-6 88.91 P-M NN11 late Miocene amplificus (6.91) (7.42) IWA U1382B- B Nicklithus B Amaurolithus spp. 10H-6 89.11 P-M NN11 late Miocene amplificus (6.91) (7.42) IWB U1328B- B Nicklithus B Amaurolithus spp. 10H-7- 90.41 P-M NN11 late Miocene amplificus (6.91) (7.42) IWA U1382B- B Nicklithus B Amaurolithus spp. 10H-7- 90.61 P-M NN11 late Miocene amplificus (6.91) (7.42) IWB Hole U1383D U1383D- NN20- middle-late B Emiliania T Pseudoemiliania 1H- 2 1.64 M 21 Pleistocene huxleyi (0.29) lacunosa (0.44) IWA U1383D- T Pseudoemiliania Tc Reticulofenestra asanoi 1H- 2 1.84 M NN19 early Pleistocene lacunosa (0.44) (0.91) IWB U1383D- T Pseudoemiliania Tc Reticulofenestra asanoi 3.34 P NN19 early Pleistocene 1H- 3 lacunosa (0.44) (0.91) U1383D- T Pseudoemiliania Tc Reticulofenestra asanoi 5.75 P-M NN19 early Pleistocene 2H- 1 lacunosa (0.44) (0.91) U1383D- Tc Reticulofenestra Bc Reticulofenestra 8.75 M-G NN19 early Pleistocene 2H- 3 asanoi (0.91) asanoi (1.14)

IWA

U1383D- Tc Reticulofenestra Bc Reticulofenestra 2H- 3 8.55 M-G NN19 early Pleistocene asanoi (0.91) asanoi (1.14) IWB U1383D- T Helicosphaera B Gephyrocapsa 11.75 M NN19 early Pleistocene 2H- 5 sellii (1.26) >5.5microns (1.62) T Discoaster U1383D- NN16- middle-late T Discoaster tamalis 16.77 M pentaradiatus 3H- 2 NN17 Pliocene (2.80) (2.39) U1383D- T Discoaster 3H- 4 19.57 P-M NN16 middle Pliocene T Sphenolithus spp. (3.54) tamalis (2.80) IWA U1383D- T Discoaster T Discoaster tamalis 22.77 M NN16 middle Pliocene 3H- 6 surculus (2.49) (2.80) U1383D- T Discoaster T Discoaster tamalis 25.58 P NN16? middle Pliocene 4H- 2 surculus (2.49) (2.80) U1383D- T Discoaster T Discoaster tamalis 27.58 M-G NN16 middle Pliocene 4H- 3 surculus (2.49) (2.80) U1383D- T Discoaster T Discoaster tamalis 29.08 M NN16 middle Pliocene 4H- 4 surculus (2.49) (2.80) U1383D- T Discoaster 30.58 M NN16 middle Pliocene T Sphenolithus spp. (3.54) 4H- 5 tamalis (2.80) T Reticulofenestra U1383D- NN13- B Ceratolithus rugosus 32.08 P-M early Pliocene pseudoumbilicus 4H- 6 15 (5.12) (3.70) U1383D- T Discoaster 33.58 P-M NN16 middle Pliocene T Sphenolithus spp. (3.54) 4H- 7 tamalis (2.80) U1383D- T Discoaster 37 P NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H- 3 tamalis (2.80) U1383D- T Discoaster 38.7 P-M NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H -4 tamalis (2.80) U1383D- T Discoaster 40 P NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H- 5 tamalis (2.80) T Reticulofenestra U1383D- NN13- B Ceratolithus rugosus 41.5 P-M early Pliocene pseudoumbilicus 5H- 6 15 (5.12) (3.70) T Reticulofenestra U1383D- NN13- B Ceratolithus rugosus 43 P early Pliocene pseudoumbilicus 5H- 7 15 (5.12) (3.70) Hole U1383E U1383E- NN19 T Calcidiscus T Discoaster surculus 1.3 P-M late Pliocene? 1H-1 (early) macintyrei (1.6) (2.49) U1383E- late Pliocene/early T Pseudoemiliania T Gephyrocapsa >5.5 2.7 M-G NN19 1H-2 Pleistocene lacunosa (0.44) microns (1.24) U1383E- late Pliocene/early T Pseudoemiliania B Gephyrocapsa 5.08 M NN19 2H-2 Pleistocene lacunosa (0.44) >5.5microns (1.62) U1383E- late Pliocene/early T Pseudoemiliania T Discoaster brouweri 8.08 P-M NN19 2H-4 Pleistocene lacunosa (0.44) (1.93) T Discoaster U1383E- NN16- middle-late T Reticulofenestra 11.08 P-M pentaradiatus 2H-6 17 Pliocene pseudoumbilicus (3.70) (2.39) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 17.58 P-M early Pliocene pseudoumbilicus 3H-4 15 (5.12) (3.70) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 20.58 P-M early Pliocene pseudoumbilicus 3H-6 15 (5.12) (3.70) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 23.26 P-M early Pliocene pseudoumbilicus 4H-2 15 (5.12) (3.70) U1383E- T Discoaster 26.26 P-M NN16 middle Pliocene T Sphenolithus spp. (3.54) 4H-4 surculus (2.49)

U1383E- T Discoaster 27.76 P NN16 middle Pliocene T Sphenolithus spp. (3.54) 4H-5 surculus (2.49) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 29.26 P-M early Pliocene pseudoumbilicus 4H-6 15 (5.12) (3.70) U1383E- T Discoaster 32.5 P-M NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H-1 tamalis (2.80) U1383E- T Discoaster 33.75 M NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H-2 tamalis (2.80) U1383E- T Discoaster 35.25 M-G NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H-3 tamalis (2.80) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 36.75 P-M early Pliocene pseudoumbilicus 5H-4 15 (5.12) (3.70) U1383E- T Discoaster 38.45 M NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H-5 tamalis (2.80) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 39.75 M early Pliocene pseudoumbilicus 5H-6 15 (5.12) (3.70) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 42.53 P early Pliocene pseudoumbilicus 6H-2 IWA 15 (5.12) (3.70) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 42.73 P early Pliocene pseudoumbilicus 6H-2 IWB 15 (5.12) (3.70) T Reticulofenestra U1383E- NN13- B Ceratolithus rugosus 48.13 P early Pliocene pseudoumbilicus 6H-6 15 (5.12) (3.70) Hole U1384A U1384A- NN20- Middle-Late T Pseudoemiliania 1.17 G modern 1H-1 21 Pleistocene lacunosa (0.44) U1384A- NN20- Middle-Late T Pseudoemiliania 2.67 G modern 1H-2 21 Pleistocene lacunosa (0.44) U1384A- T Helicosphaera B Gephyrocapsa 6.54 M NN19 early Pleistocene 2H-2 sellii (1.26) >5.5microns (1.62) U1384A- T Pseudoemiliania T Gephyrocapsa 9.54 G NN19 early Pleistocene 2H-4 IWA lacunosa (0.44) >5.5microns (1.24) U1384A- T Helicosphaera B Gephyrocapsa 12.54 M-G NN19 early Pleistocene 2H-6 sellii (1.26) >5.5microns (1.62) U1384A- NN16- middle Pliocene- T Pseudoemiliania T Discoaster tamalis 15.78 M 3H-2 IWB 19 early Pleistocene lacunosa (0.44) (2.80) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 20.28 M early Pliocene pseudoumbilicus 3H-5 15 (5.12) (3.70) U1384A- T Discoaster T Discoaster tamalis 25.29 M NN16 middle Pliocene 4H-2 IWB surculus (2.49) (2.80) U1384A- T Discoaster T Discoaster tamalis 28.29 M NN16 middle Pliocene 4H-4 IWB surculus (2.49) (2.80) U1384A- T Discoaster T Discoaster tamalis 31.29 M NN16 middle Pliocene 4H-6 surculus (2.49) (2.80) U1384A- T Discoaster 34.54 M-G NN16 middle Pliocene T Sphenolithus spp. (3.54) 5H-2 tamalis (2.80) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 40.54 M early Pliocene pseudoumbilicus 5H-6 15 (5.12) (3.70) U1384A- NN13/ B Ceratolithus B Ceratolithus rugosus 45.4 M early Pliocene 6H-2 IWB 15 rugosus (5.12) (5.12) U1384A- T Discoaster T Reticulofenestra 48.4 M NN16 middle Pliocene 6H-4 IWB tamalis (2.80) pseudoumbilicus (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 49.9 M early Pliocene pseudoumbilicus 6H-5 IWB 15 (5.12) (3.70) U1384A- NN13/ T Reticulofenestra B Ceratolithus rugosus 51.2 M early Pliocene 6H-6 IWA 15 pseudoumbilicus (5.12)

(3.70)

T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 54.58 M early Pliocene pseudoumbilicus 7H-2 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 56.08 M early Pliocene pseudoumbilicus 7H-3 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 57.58 P early Pliocene pseudoumbilicus 7H-4 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 58.88 M early Pliocene pseudoumbilicus 7H-5 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 59.98 M early Pliocene pseudoumbilicus 7H-6 IWA 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 67.97 M early Pliocene pseudoumbilicus 8H-5 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 69.47 M early Pliocene pseudoumbilicus 8H-6 15 (5.12) (3.70) T Reticulofenestra U1384A- NN13/ B Ceratolithus rugosus 70.77 M early Pliocene pseudoumbilicus 8H-7 IWA 15 (5.12) (3.70) U1384A- T Nicklithus B Nicklithus amplificus 75.91 M NN11 late Miocene 9H-4 amplificus (5.94) (6.91) U1384A- late Miocene/early T Ceratolithus B Ceratolithus 77.41 M NN12 9H-5 Pliocene acutus (5.04) larrymeyeri (5.34) T Discoaster U1384A- T Nicklithus amplificus 78.33 M NN11 late Miocene quinqueramus 9H-6 (5.94) (5.59) T Discoaster U1384A- T Nicklithus amplificus 79.83 M NN11 late Miocene quinqueramus 9H-7 IWB (5.94) (5.59) T Discoaster U1384A- T Nicklithus amplificus 85.5 M NN11 late Miocene quinqueramus 10H-4 (5.94) (5.59) U1384A- NN11- late Miocene/early 87 P — — 10H-5 12 Pliocene U1384A- T Nicklithus B Nicklithus amplificus 88.5 M NN11 late Miocene 10H-6 amplificus (5.94) (6.91) U1384A- T Nicklithus B Nicklithus amplificus 10H-7 90 M NN11 late Miocene amplificus (5.94) (6.91) IWB U1384A- T Nicklithus B Nicklithus amplificus 11H-2- 91.8 P NN11 late Miocene amplificus (5.94) (6.91) IWB U1384A- B Amaurolithus B Discoaster berggrenii 11H-3 93.1 M NN11 late Miocene spp. (7.42) (8.29) IWA U1384A- B Amaurolithus B Discoaster berggrenii 11H-3 93.3 M NN11 late Miocene spp. (7.42) (8.29) IWB U1384A- 11H-4- 94.5 VP — — — — IWA U1384A- 11H-4- 94.7 VP — — — — IWB aAbbreviations: G, good; M, moderate P, poor; VP, very poor

Table S2-2. North Pond total sedimentary P and accumulation rates P acc. ratea Depth Sed. ratea Sample ID TPa (µmol g-1) DBDa,b (g cm-3) (µmol cm-2 Age (Ma) (mbsf) (cm kyr-1) kyr-1)

Hole U1382B

1H-2 0.75 12.9 0.86 1.26 14.0 0.44 2H-5 – 2H-6 12.3 14.5 0.97 1.26 17.6 1.26 3H-4 – 3H-5 21.0 10.3 1.07 1.26 13.9 1.60 3H-5 – 6H-7 36.6 10.3 1.04 1.26 13.5 2.80 8H-6 – 8H-7 71.2 13.0 0.93 1.26 15.1 3.70 9H-6 – 9H-7 80.8 15.8 0.97 1.26 19.3 5.12 9H-6 – 10-2 81.4 16.6 1.00 1.26 21.0 5.59 9H-6 – 9H-7 80.8 15.8 0.97 1.26 19.3 5.94 10H-4 – 10H-5 86.8 33.8 1.06 1.26 44.9 6.91

Hole U1383D 1H-2A – 1H-2B 1.74 13.9 0.83 1.10 12.6 0.44 2H-1 – 2H-3 7.25 14.5 0.96 1.10 15.3 0.91 2H-3 – 2H-5 10.2 13.1 0.95 1.10 13.8 1.14 2H-3 – 2H-5 10.2 13.1 0.95 1.10 13.8 1.26 2H-5 – 3H-2 14.3 11.6 0.93 1.10 11.9 1.62 4H-4 – 4H-5 29.8 12.5 0.97 1.10 13.3 2.80 5H-5 – 5H-6 40.8 13.0 1.02 1.10 14.6 3.54 5H-5 – 5H-6 40.8 13.0 1.02 1.10 14.6 3.70 Hole U1383E

1H-2 1.35 12.6 0.79 1.06 10.6 0.44 2H-4 – 2H-6 9.58 15.5 0.93 1.06 15.3 1.93 2H-6 – 4H-4 18.7 12.3 1.02 1.06 13.2 2.49 5H-5 – 5H-6 39.1 13.8 0.98 1.06 14.4 3.54 5H-5 – 5H-6 39.1 13.8 0.98 1.06 14.4 3.70 Hole U1384A

1H-2 – 2H-2 4.61 10.9 0.94 1.25 12.7 0.44 1H-2 – 2H-2 4.61 10.9 0.94 1.25 12.7 1.26 2H-6 – 3H-2 14.2 15.1 1.02 1.25 19.2 1.62 2H-6 – 4H-6 18.9 13.5 1.03 1.25 17.3 2.49 4H-6 – 5H-2 32.9 12.6 0.97 1.25 15.3 2.80 8H-7 – 9H-4 73.3 12.9 1.06 1.25 17.1 5.12 9H-5 – 9H-6 77.9 16.0 1.02 1.25 20.2 5.34 9H-5 – 9H-6 77.9 16.0 1.02 1.25 20.2 5.59 10H-4 – 10H-6 87.0 26.3 1.06 1.25 34.9 5.94 11H-2 – 11H-3 92.5 54.6 0.72 1.25 48.8 7.42 aAbbreviations: TP, total P; DBD, dry bulk density; sed. rate, sediment accumulation rate; P acc. rate, P accumulation rate bData from Expedition 336 scientists [2012]

Table S2-3. Grizzly Bare total sedimentary P and accumulation rates

Average TPa Average Sed. ratea P acc. ratea (µmol Deposit type (µmol g-1) DBDa,b (g cm-3) (cm kyr-1) cm-2 kyr-1)

c Hemipelagic mud 20.8 1.14 10.19 242 d Sandy turbidite 20.8 1.14 11.83 281 d Sandy turbidite 20.8 1.14 55.88 1330 aAbbreviations: TP, total P; DBD, dry bulk density; sed. rate, sediment accumulation rate; P acc. rate, P accumulation rate bDBD measurements conducted by IODP Expedition 327 shipboard scientists [Expedition 327 scientists, 2011] cData from [Underwood et al., 2005] dData from [Su et al., 2000]

Chapter Three

POTENTIAL PHOSPHORUS UPTAKE MECHANISMS IN THE DEEP

SEDIMENTARY BIOSPHERE

In prep: Defforey, D., Tully, B.J., Sylvan, J.B., Cade-Menun, B.J., Reese, B.K. and Paytan, A. (2016). Potential phosphorus uptake mechanisms in the deep sedimentary biosphere. Environ. Sci. Technol. (in prep)

89 Abstract

In this study, we explore potential microbial P uptake mechanisms in oligotrophic marine sediments beneath the North Atlantic Gyre and their effects on the relative distribution of organic P compounds as a function of burial depth and changing redox conditions. Metagenomic analyses were used to investigate the presence of microbial functional genes pertaining to P uptake and metabolism. These analyses were coupled with solution 31P nuclear magnetic resonance spectroscopy (31P NMR) to shed light on possible P substrates for the deep biosphere in these organic matter depleted sediments. This is the first time, to our knowledge, that solution 31P NMR has been used to explore changes in organic P compounds in oligotrophic marine sediments at depths down to 57 m below seafloor. We identified a wide variety of organic P compounds, including inositol hexakisphosphate stereoisomers, nucleic acids, sugar phosphates, phosphonates and phospholipids, in addition to pyrophosphate and polyphosphates. Some of the genes identified include genes related to phosphate transport, phosphonate and polyphosphate metabolism, as well as phosphite uptake.

Results from our study suggest that the deep sedimentary biosphere may have adapted to take advantage of a wide array of P substrates and could play a role in the gradual breakdown of inositol and sugar phosphates, as well as reduced P compounds and polyphosphates.

90 1 Introduction

Phosphorus (P) is a key macronutrient for life involved in the structure of nucleic acids in the form of a phosphate–ester backbone, and in the transmission of chemical energy by the adenosine triphosphate (ATP) molecule. P is also a structural constituent in many cell components such as phosphoproteins, and phospholipids in cell membranes. In addition, P is involved in many cellular processes through the mechanism of phosphorylation, which alters the function and activity of proteins.

Hence, P is required to sustain life, including microbial life in the deep subseafloor environment. This environment is estimated to comprise as many prokaryotic cells as the marine water column and soils, and includes significant prokaryotic populations at depths greater than two kilometers [Roussel et al., 2008; Kallmeyer et al., 2012;

Inagaki et al., 2015]. Recent estimates, based on cell counts from both ocean margins and oligotrophic ocean gyres sediments, suggest that the deep sedimentary subseafloor biosphere could contain 0.18-3.6% of Earth’s total biomass and an equal amount of microbial cells as the marine water column (Kallmeyer et al., 2012). The existence and activity of deep subseafloor sediment microbial populations have profound implications on global biogeochemical cycles and our understanding of the limits of life. However, there is a substantial knowledge gap regarding the nature of P compounds sustaining life in oligotrophic open ocean sediments, and how microbial activity impacts the relative distribution of P compounds in general, and organic P compounds in particular in marine sediments. In addition, little is know about the impact of the deep biosphere on sedimentary P cycling and P diagenetic processes.

91 This is largely because of analytical difficulties associated with studying P cycling and particularly organic P in deep ocean sediments that contain low concentrations of organic matter [Ruttenberg, 2014]. Therefore, a better understanding of the mechanisms by which deep subseafloor microbes obtain this essential nutrient is needed.

This study’s field site is located at North Pond, an approximately 8 × 15 km sediment-filled basin located on the western flank of the Mid-Atlantic ridge sampled during IODP Expedition 336 [Expedition 336 Scientists, 2012a]. Such sediment ponds surrounded by high relief topography are common to the western flank of the

Mid-Atlantic Ridge, and thought to be representative of slow-spreading ridge flank environments [Becker et al., 2001]. Sediments at North Pond are predominantly nannofossil ooze with layers of coarse foraminiferal sand and occasional pebble-sized clasts of basalt, serpentinite, gabbroic rocks, and bivalve debris [Expedition 336

Scientists, 2012a]. Organic carbon and nitrogen contents in those sediments are low

[Ziebis et al., 2012], and organic P was detected at low concentrations throughout the sediment column using sequential sediment extractions [Defforey and Paytan, 2015].

In addition, oxygen penetrates deep in the sediment column, and has a “C” shaped depth profile due to oxygen fluxes from oxygenated seawater at the sediment- seawater interface and oxygenated fluids in the underlying basalt crust, with the middle portion of the cores being suboxic to anoxic [Orcutt et al., 2013]. Deep sea,

92 carbon-depleted sediments such as the ones sampled in this study represent approximately 70% of the seafloor [Mewes et al., 2016].

In this study, we present metagenomic data from a sediment sample collected at 59 m below seafloor (mbsf), as well as solution 31P nuclear magnetic resonance spectroscopy spectra (31P NMR) from two depths in the sediment column (3.2 and

56.7 mbsf). The combination of these data yields insights into putative P uptake mechanisms and P substrates available for deep subseafloor microorganisms. This work also helps improve our knowledge of the role deep subseafloor bacteria play in

P cycling and our understanding of the chemical composition of organic P in organic- lean oligotrophic sediments.

2 Materials and Methods

2.1 Sample collection and storage

The sediment samples used in this study were collected from Site U1382 (Hole

U1382B) at North Pond using the advanced piston corer during IODP Expedition 336

(Figure 1). Site U1382 is located in the southeastern part of North Pond, where sediments are approximately 90 m thick [Expedition 336 scientists, 2012d]. The sample used for metagenomic analyses was retrieved from the suboxic part of the sediment column (at 59.1 mbsf, 1 µM dissolved oxygen) and the samples used for

31PNMR were retrieved from oxic (30 µM DO) and suboxic (1 µM DO) portions of the sediment column respectively (at 3.2 mbsf and 56.7 mbsf) [Orcutt et al., 2013].

93 Fluorescent microspheres (Fluoresbrite Carboxylate Microspheres; Polysciences, Inc.,

15700) were used in every core recovered to monitor any potential contamination

[Expedition 336 scientists, 2012a]. Upon recovery, whole-round sediment cores, including sample U1382B 7H-5, which we used for metagenomic analyses, were sectioned on the catwalk, capped and stored at –80oC in sterile bags [Expedition 336 scientists, 2012b]. The sediment samples used for 31P NMR analyses were squeezed- cakes stored at –80oC after porewater collection immediately following core retrieval.

2.2 DNA extraction

DNA was extracted using the method described in [Mills et al., 2012] as modified by

B. K. Reese for nucleic acids in low biomass open ocean sediments. The interior of the whole-round core for sample U1382B 7H-5 was subsampled using sterile techniques and was divided into 25 splits, each weighing ~0.5 g. Initial cell lysis was achieved using five cycles of freeze (liquid nitrogen), thaw (55oC water bath) and vortex steps while stabilizing nucleic acids in a Tris–EDTA–glucose buffer. This step was followed by the addition of lysozyme to the buffer and incubation at 30oC for 10 minutes. Samples were then treated twice with buffered phenol, chloroform and isoamyl alcohol (25:24:1; pH 8.0), and sodium dodecyl sulfate to dissolve the cell membrane and solubilize both high and low molecular weight proteins. Nucleic acids within the aqueous layer above the phenol–chloroform layer were then precipitated in an ethanol and sodium acetate solution. The ethanol solution was decanted following centrifugation of samples at 4°C. Lastly, DNA pellets were resuspended in sterile

94 water, combined into one sample and the concentration of DNA was quantified using a Qubit fluorometer (Life Technologies). Extraction blanks were included with each sample extraction and yielded no measurable DNA.

2.3 Bioinformatic analyses

Raw reads were quality controlled using cutadapt (v1.7.1) and Trimmomatic (v0.33).

The QCed sequences were searched for SSU rRNA fragments using Meta-RNA

(v.H3). SSU rRNA fragments were assembled using EMIRGE (v.1.3) utilizing

SILVA SSU as the reference database using the SINA web portal aligner [Ludwig et al., 2004; Capella-Gutiérrez et al., 2009; Pruesse et al., 2012]. 23 full-length SSU rRNAs were reconstructed for sample U1382B 7H-5. Taxonomy was assigned to these assemblies using the mothur (v.1.3.7) commands align.seqs and classify.seqs and SILVA SSU as the reference database. The quality checked (QCed) sequence library was aligned to its assembled SSU rRNAs using Bowtie2 (v2.2.5). Based on the total number of QCed sequences that aligned to the SSU rRNAs, a length- normalized relative abundance value was determined. Results showed that the sample contained some degree of human-related contamination, both from human DNA and human microbiome DNA. In order to verify that the functional genes related to P metabolism were not from human microbiome DNA sequences, we constructed a 16S rRNA phylogenetic tree from the metagenome to help determine the origin of microbial taxa (Supplementary Fig. S1). Those most closely related to environmental taxa were considered valid, while those similar to known contaminants, such as

95 common components of the human skin microbiome, were considered contaminants.

We also excluded any sequences from known human microbiome

(Propionobacterium, Microbacteriaceae), as well as sequences without high-level taxonomic assignment from the Proteobacteria (that could include Enterobacter). The

QCed sequence library was assembled using IDBA-UD (v.1.1.1), but the overall quality of these assemblies was low (contigs: 49614 n50: 467 max: 48419 mean:

461). A second round of assembly using all of the contigs from the IDBA-UD output was performed using the Geneious De Novo assembler (v.6.3.8) [Kearse et al., 2012].

This step generated a total of approximately 130 contigs >2.5kb in length. This number was too small for genome binning analysis. To identify genes within the assembled dataset related to phosphorous metabolism, all putative coding DNA sequences (CDSs) were determined using Prodigal (v2.6.3), using the metagenome option (-p meta).

2.4 16S rRNA phylogenetic tree

Full-length 16S rRNA sequences were generated using EMIRGE (as described above) and were greater than 1,000 bp in length, with the exception of ID 973 due to poor resolution within the sequence. The sequences were aligned to the SILVA SSU reference database using the SINA web portal aligner (https://www.arb- silva.de/aligner/) [Ludwig et al., 2004; Capella-Gutiérrez et al., 2009; Pruesse et al.,

2012]. These alignments were loaded in to ARB (v6.0.3), manually assessed and

96 added to the non-redundant 16S rRNA gene database (SSURef123 NR99) using ARB

Parsimony (Quick) tool (parameters: default). We identified a selection of the nearest neighbors to the sediment metagenome sequences to be used to construct a 16S rRNA phylogenetic tree. We also identified additional sequences related to the environmental 16S rRNA using a BLAST search against the NCBI RefSeq and non- redundant databases. Environment and reference 16S rRNA sequences were aligned using MUSCLE (v3.8.31; parameters: -maxiters 8), processed by the automated trimming program trimAL (v1.2rev59; parameters: -automated1), and aligned for a second time using MUSCLE (parameters: -maxiters 8). The alignment was used to construct an approximate maximum-likelihood phylogenetic tree using FastTree

(v2.1.3; parameters: -nt -gtr -gamma) [Price et al., 2010].

2.5 Solution 31P NMR extraction

Two sediment samples, U1382B 1H-3 (3.2 mbsf) and 7H-5 (56.7 mbsf), were processed following the method described in Defforey et al. [2016, in review]. This method quantitatively removes orthophosphate prior to the alkaline extraction for solution 31P NMR to amplify the signal of low abundance P compounds, including organic P, relative to that of orthophosphate so that these P forms in oligotrophic sediments may be resolved without the need for extremely long NMR experiments.

Briefly, each freeze-dried, ground and sieved sample (<125 µm) was divided into 8 splits (1.5 g each). Sample splits were extracted in a citrate–dithionite–bicarbonate buffer to solubilize P bound to iron (Fe) oxyhydroxides (8 h; pH 7.6) and the

97 sediment residues washed with ultrapure water (18.2 MΩ·cm deionized water; 2 h).

The samples were then extracted in a sodium acetate buffer (6 h; pH 4.0) to solubilize

P bound to carbonate fluorapatite, and washed with ultrapure twice (2 h each). After this step, the 8 splits for each sample were combined, and extracted in 0.25 M NaOH

+ 0.05 M Na2EDTA over the course of 6 h. The alkaline extracts were frozen and lyophilized, and then redissolved in 500 µL D2O, H2O, NaOH–EDTA and 10 M

NaOH prior to solution 31P NMR analysis. Sediment residues were ashed at 550°C and extracted in 0.5 M sulfuric acid (16 h) to determine residual P content.

2.6 Solution 31P NMR analysis and post-experimental processing

Spectra were acquired immediately following sample preparation on a Varian Unity

INOVA 600 MHz spectrometer equipped with a 10 mm broadband probe [operated by the Stanford Magnetic Resonance Laboratory at Stanford University]. The analytical parameters used were: 20oC, 90o pulse, 0.48 s acquisition time, 4.52 s delay time, 5600 scans (8 h experiments), no spin and an external H3PO4 standard. No proton decoupling was used out of concern for sample degradation [Cade-Menun and

Liu, 2014]. We used P to Fe and manganese (Mn) ratios as a proxy for spin-lattice relaxation times (T1) to ensure adequate delays between pulses and thus quantitative spectra [McDowell et al., 2006]. We used 5 s recycle delays, which correspond to three to five times the calculated T1 values. Peak identification was based on spiking with known compounds (2-aminoethylphosphonic acid, α– and β–glycerophosphate, glucose-6-phosphate) and on literature values [Cade-Menun, 2015]. Monoester 1

98 refers to unidentified orthophosphate monoesters located between 7–6.1 ppm, while monoesters 2 and 3 include peaks between 5.7–3.6 ppm and 4.0–2.5 ppm respectively

[Cade-Menun et al., 2015]. We processed 31P NMR data using the NMR Utility

Transform software (NUTS, Acorn NMR) and calculated peak areas by integration of spectra processed with a 7 Hz line broadening for the full spectrum and 2 Hz line broadening for the monoester region following baseline correction, peak picking and phasing. We accepted peaks that (1) represented at least 1% of the tallest peak in the total integrated area, (2) were identified by the NUTS software and (3) were confirmed as signal by visual inspection.

2.7 Supernatant P analyses

We measured total P concentrations in sediment extracts using inductively coupled plasma optical emission spectroscopy (ICP-OES) [Perkin-Elmer Optima 4300 DV

Inductively Coupled Plasma Optical Emission Spectrometer operated by the

University of California, Santa Cruz]. Standards were prepared with the same solutions as those used for the extraction procedure and ranged from 0 – 320 µM P.

The detection limit for P on this instrument for both wavelengths is 0.4 µM. We measured molybdate reactive P concentrations (MRP, which includes primarily free orthophosphate) on a QuikChem 8000 automated ion analyzer. Standards were prepared with the same solutions used for the extraction step to minimize matrix effects on P measurements, and ranged from 0 – 30 µM P. The detection limit for P on this instrument is 0.2 µM. We derived molybdate unreactive P concentrations

99 (MUP, which includes primarily organic P and polyphosphates) by subtracting MRP from total P concentrations.

3 Results and Discussion

A target subset of P-related genes were identified in sample U1382B 7H-5 (59.1 mbsf), consisting of 23 genes of interest including phosphate utilization (transport, pstABS; metabolism, phoABDEHR), phosphonate utilization (transport and metabolism; phnACDGJ), phosphite utilization (ptxABCD), glycerol 3-phosphate transport (ugpBC), polyphosphate metabolism (ppKX), and phosphatase activity

(appAB) (Table 1). This set of genes could be split into two groups. Group 1, consisting of pstABS, ppX, ugpBC, ptxA, phoDR, were present in the FOAM database (accessed April 2016), a curated set of hidden Markov models (HMMs) generated using the KEGG ontology database that could be used to search conserved motifs within the environmental sequences. For FOAM searchable genes of interest

(group 1), environmental sequences (excluding phoR and upgC) were searched using hmmsearch (v3.1b2; parameters, --cut_tc --notextw) with a bit score cutoff of 72

(approximately equivalent to an e-value of 1×10-20). Gene sequences for phoR and upgC represented protein subunits with generally highly conserved regions: sensor histidine kinases and ATP-binding motifs for ABC-type transporters, respectively. As such, these matches could not be trusted at the same cutoff enforced for the other genes within group 1, and a more stringent cutoff of 105 bit score (approximate e-

100 value equivalent 1×10-50) was used. The results from the FOAM database identified the presence of ppX (exopolyphosphatase) and pstAB, the ATP-binding and permease subunits of the phosphate ABC-type transporter system. However, the crucial pstS, solute-binding subunit, was not identified. Furthermore, even with the more stringent cutoff values, 23 and 38 copies of upgC and phoR were identified. This large number suggests that even at this cutoff value, a more diverse group of these gene families were identified. The remaining set of genes of interest (group 2), including ptxABCD, appA, ppK, phoABED and phnACD were used to search the remaining putative CDS using BLASTP (v 2.2.30+, parameters, e-value of 1×10-20, max_target_seqs 1). A total of 187 matches to the environmental sequences were identified. However, the problematic nature of the conserved region within the ATP-binding subunit of the

ABC-type transporters resulted in 107 matches to environmental sequences. These were deemed uninformative and were excluded from further analysis. Matches were identified for appAB, phoD, phnADGJ, ptxCD, ppK. A large portion of the BLAST matches (after ATP-binding subunits) belonged to phoB (n = 37), a phosphate regulon response regulator, but this protein family is diverse and multiple types of regulon genes may be identified in this manner. Lastly, for each of the genes identified as related to P uptake and metabolism (both FOAM and BLAST databases), a putative taxonomy was assigned. Putative P-related environmental genes were queried using

BLASTP (parameters, e-value 1×10-5, -max_target_seqs 5 -outfmt 5) against the

GenBank RefSeq (v75) database. The MEGAN (v4) Last Common Ancestor (LCA) algorithm was used to assign a putative taxonomy based on the top five BLAST

101 matches (Table 1). Most P related gene sequences belonged to Alphaproteobacteria, or to a combination of Alphaproteobacteria, Betaproteobacteria, Actinobacteria and unclassified microorganisms (Table 1). The metagenomic analyses performed in this study only yield information regarding the presence of functional genes pertaining to

P metabolism, and these may represent DNA from active and inactive or dormant microorganisms. Indeed, most phylogenetic groups identified in North Pond sediments have been shown to be inactive [B. K. Reese, personal communication].

However, active Alphaproteobacteria (mainly from the Caulobacteraceae family and

Brevundimonas genus) have been found throughout the sediment column at Hole

U1382B, as well as some active Betaproteobacteria (Achromobacter genus) and

Gammaproteobacteria (Pseudomonas genus, and the genera Stentrophomonas and

Methylophaga) [B. K. Reese, personal communication]. This suggests that the functional genes reported in our study are present at depths where microorganisms are active and utilizing P and may thus be impacting sedimentary organic matter remineralization through their metabolic activity.

Solution 31P NMR spectra for both samples are shown in Figure 2. The 2–step pretreatment preceding the alkaline extraction removed 95% of orthophosphate in sample U1382B 1H-3, and 98% in sample U1382B 7H-5 with no measurable MUP removal (Table 3; Figure 3). For both samples, the 5 s recycle delays were long enough to produce quantitative spectra (Table 4). Metal content was similar in both

NMR extracts, but P content was lower for the deeper sample (Table 4). The

102 shallower, oxic sample (U1382B 1H-3, 3.2 mbsf, 30 µM DO) contains a wider variety of P compounds than the deeper, suboxic sample (U1382B 7H-5, 56.7 mbsf,

~1 µM DO). The most abundant P compounds in both samples identified using solution 31P NMR after the 2–step pretreatment are inorganic orthophosphate and pyrophosphate (Table 2; Figure 2). The latter is a byproduct of the biosynthesis of a number of macromolecules, including nucleic acids, amino acids, coenzymes, nucleotides and activated precursors to proteins and polysaccharides [Heinonen,

2001]. We also measured small amounts of organic P compounds, including phosphonates (reduced organic P compounds with a direct C–P bond), orthophosphate monoesters, diesters (DNA and possibly some phospholipids) and polyphosphates in both samples (Table 2; Figure 2). Orthophosphate monoesters are the most abundant forms of organic P in both of our samples, which is consistent with numerous studies that have shown them to generally be the dominant group of organic P compounds in both marine and terrestrial environments [Ingall et al., 1990;

Clark et al., 1998, 1999; Carman et al., 2000; Kolowith et al., 2001; Paytan et al.,

2003; Turner et al., 2005; Sannigrahi and Ingall, 2005; Sannigrahi et al., 2006]. In the shallower sample, we were able to resolve individual peaks in the orthophosphate monoester region and identify a number of different monoester compounds (Table 2).

Among them, the most abundant type of monoesters identified was inositol hexakisphosphate (IHP) stereoisomers: myo–, scyllo–, neo– and D-chiro–IHP.

Inositol phosphates form the dominant class of organic P compounds in soils, and abundant in aquatic systems [Turner et al., 2002]. Of all IHP stereoisomers, myo–IHP

103 is the most abundant, but scyllo–, D–chiro– and neo–IHP have also been measured in soils and marine sediments [White and Miller, 1976; Turner et al., 2012]. Also present are glucose-6-phosphate, choline phosphate, α– and β–glycerophosphate.

However, we were unable to resolve individual orthophosphate monoester peaks from background noise in the deeper sample (56.7 mbsf) due to low abundance of these compounds in the NMR extracts.

The combined use of metagenomic analyses and solution 31P NMR spectroscopy yields valuable information regarding possible P substrates available to deep sedimentary subseafloor microorganisms, as well as potential microbial P uptake mechanisms. While we do not have a metagenome for the shallower sediments at

Hole U1382B, the decrease in abundance of organic and biogenic P (pyrophosphate, polyphosphates) suggest that deep sedimentary subseafloor microorganisms are utilizing those compounds as a P source, and possibly as an organic C source in this

C-depleted environment [Heath, 2005; Ziebis et al., 2012] (Table 2).

Myo-IHP and glucose 6-phosphate, both present in the shallower sample, are thought to adsorb on the same sites of Fe and aluminum (Al) oxides as phosphate anions since adsorption occurs through the phosphate groups via ligand exchange with surrounding porewater and hydroxide groups on mineral surfaces [Goldberg and

Sposito, 1985; Baldwin et al., 2002; Celi and Barberis, 2005]. Little is known regarding possible differences in the behavior of other stereoisomeric forms of IHP,

104 but they may adsorb on oxide surfaces in a similar manner as myo-IHP. Once sorbed on short-range ordered aluminum precipitates, inositol IHP and glucose-6-phosphate have been shown to be poorly available to microorganisms [Shang et al., 1996], which would allow them to persist at depth in marine sediments. However, surface properties of oxides are modified when associated with other minerals such as clay minerals, which affect their interactions with organic P and may weaken the complex formed with oxides [Celi et al., 2003], thus increasing their bioavailability. In addition to sorption on Fe and Al oxides, myo-IHP can form soluble complexes with calcium in calcareous sediments, which may enhance the interactions between myo-

IHP and calcium-bearing minerals and further stabilize it [Celi and Barberis, 2005].

The bioavailability of organic phosphorus compounds is primarily dependent on the stability of the phosphate surface complexes rather than by the total amount of phosphate adsorbed [Shang et al., 1996]. Varying redox conditions have been shown to strongly regulate IHP remineralization, and lead to enhanced IHP decomposition under anaerobic conditions [Suzumura and Kamatani, 1995]. The observed decrease in orthophosphate monoester abundance with depth could be due to an increase in bioavailability of those compounds resulting from the transition to suboxic conditions in the middle parts of the sediment column [Zhang et al., 1994] (Table 2, Figure 2).

Indeed, bacteria at this depth possess genes encoding for phosphomonoesterases,

(appA, appB, phoD) [Vershinina and Znamenskaya, 2002] and could utilize those deeply buried P compounds as they become more susceptible to enzymatic hydrolysis.

105

Orthophosphate diesters are also present at both depths in the form of DNA in the shallow sample, and possibly DNA and phospholipids in the deeper sample (Table 2,

Figure 2). The sorption of nucleic acids on mineral surfaces is affected by the molecular weight of DNA, the presence of montmorillonite, pH and redox conditions

[Greaves and Wilson, 1969; Celi and Barberis, 2005]. The complexed nucleic acid forms with montmorillonite are weaker than those with myo–IHP due to the presence of a single hydroxide group available for ligand exchange [Greaves and Wilson,

1969]. This makes nucleic acids more vulnerable to microbial enzymatic hydrolysis

[Greaves and Wilson, 1970]. The decrease in orthophosphate diester abundance with depth and the presence of genes such as phoD and ugpC suggest putative mechanisms for the uptake of orthophosphate diesters (phoD) and glycerophosphoryl diesters

(ugpC) at Site U1382 [Wanner, 1993; Vershinina and Znamenskaya, 2002].

Other potential sources of P, and possibly C, to the deep biosphere are phosphonates, which we measured both in shallow and deeper sediments (Table 2, Figure 2). While previously thought to be refractory, these compounds have been shown to be as bioavailable as phosphate esters [Laarkamp, 2000], especially under anoxic conditions [Benitez-Nelson et al., 2004] and represent a potential source of both P and organic C to the deep biosphere. Indeed, a number of genes pertaining to phosphonate utilization were identified in our sample: phnA (phosphonoacetate hydrolase), phnD

(phosphonate-binding periplasmic protein), phnJ (component of C-P lyase complex)

106 and phnG (component of C-P lyase complex) (Table 1). The observed decrease in phosphonate abundance (from 6.7 nmol g-1 to 2.2 nmol g-1), the high ester to phosphonate ratios, the presence of genes related to phosphonate utilization and redox conditions shown to enhance phosphonate remineralization (i.e. reducing conditions) suggest that phosphonates represent a likely substrate for deep subseafloor microorganisms at that depth in the sediment column.

Small amounts of polyphosphates were measured in both spectra, and genes for exopolyphosphatase (ppX) were present in the metagenome (Table 1, Figure 2). This enzyme catalyzes the hydrolysis of the terminal phosphate group in polyphosphates

[Jones et al., 2016]. We also identified gene sequences for polyphosphate kinase

(ppK), which catalyzes the reversible synthesis of polyphosphate from nucleoside triphosphates [Jones et al., 2016]. Polyphosphates are synthesized by all cells, and play an important role in nutrient storage, regulatory functions and metal chelation

[Rao et al., 2009]. They also play a key role in nucleating the growth of authigenic carbonate fluorapatite in marine sediments, which has profound implications for reactive P burial [Diaz et al., 2008].

An unexpected result from the metagenomic analyses was the presence of two genes related to phosphite metabolism: ptxC and ptxD. Phosphite utilization genes have been identified in diverse marine microorganisms, and their abundance is higher in marine environments with low P [Martínez et al., 2012]. Furthermore, a recent study

107 found that phosphite and phosphonate production in surface waters could contribute to a global P redox cycle [Van Mooy et al., 2015]. While we did not measure phosphite in our samples, and, to our knowledge, phosphite has not been measured in open ocean sediments, the presence of reduced organic P in the form of phosphonates and genes related to phosphite utilization raises the possibility of this compound potentially playing a role in P sediment cycling.

In this study, we present utilized a combination of metagenomics analysis and solution 31P-NMR for studying putative P uptake mechanisms and metabolism, as well as organic and biogenic P substrates available for deep subseafloor microorganisms. We demonstrated that diverse organic P compounds persist at depth in sediments underlying oligotrophic marine settings, and that changes in redox conditions may play an important role in increasing their bioavailability. We identified a wide variety of potential P sources to the deep biosphere, as well as putative enzymatic pathways for P uptake and utilization. While this study only provides potential P uptake mechanisms, it sets the stage for future work to investigate the expression of functional genes pertaining to P uptake and metabolism using RNA-based analyses and raises interesting questions about microbial transformations of sedimentary organic and biogenic P.

108 Acknowledgements

We thank the Integrated Ocean Drilling Program and the shipboard scientists and technical staff involved in IODP expedition 336 who collected the sediment samples used in this study. Funding was provided by a graduate student fellowship from the

Center for Dark Energy Biosphere Investigations (NSF C-DEBI) to D.D. (award #

157598), a student research grant from the Geological Society of America to D.D., a

C-DEBI Research and Travel Exchange Program grant to D.D., a research grant from

C-DEBI to A.P. (award # 156246) and sequencing grant to A.P., D.D., J.S. and

B.K.R. from the Census of Deep Life. We also thank Dr. Corey Liu, Rob Franks,

Michael Kong and Richard Kevorkian for their assistance with this project.

4 References

Baldwin, D. S., A. M. Mitchell, and J. M. Olley (2002), Pollutant-sediment interactions: sorption, reactivity and transport of phosphorus, in Agriculture, Hydrology and Water Quality, edited by P. M. Haygarth and S. C. Jarvis, pp. 265–280, CABI Publ., Wallingford, UK.

Benitez-Nelson, C. R., L. O’Neill, L. C. Kolowith, P. Pellechia, and R. Thunell (2004), Phosphonates and particulate organic phosphorus cycling in an anoxic marine basin, Limnol. Oceanogr., 49(5), 1593–1604, doi:10.4319/lo.2004.49.5.1593.

Cade-Menun, B. J. (2015), Improved peak identification in 31P-NMR spectra of environmental samples with a standardized method and peak library, Geoderma, 257-258, 102–114, doi:10.1016/j.geoderma.2014.12.016.

109 Cade-Menun, B. J., and C. W. Liu (2014), Solution phosphorus-31 nuclear magnetic resonance spectroscopy of soils from 2005 to 2013: a review of sample preparation and experimental parameters, Soil Sci. Soc. Am. J., 78(1), 19, doi:10.2136/sssaj2013.05.0187dgs.

Cade-Menun, B. J., Z. He, and Z. Dou (2015), Comparison of phosphorus forms in three extracts of dairy feces by solution 31P NMR analysis, Commun. Soil Sci. Plant Anal., 46(13), 1698–1712, doi:10.1080/00103624.2015.1047512.

Capella-Gutiérrez, S., J. M. Silla-Martínez, and T. Gabaldón (2009), trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses, Bioinformatics, 25(15), 1972–1973, doi:10.1093/bioinformatics/btp348.

Carman, R., G. Edlund, and C. Damberg (2000), Distribution of organic and inorganic phosphorus compounds in marine and lacustrine sediments: a 31P NMR study, Chem. Geol., 163(1), 101–114, doi:10.1016/S0009- 2541(99)00098-4.

Celi, L., and E. Barberis (2005), Abiotic stabilization of organic phosphorus in the environment, in Organic phosphorus in the environment, edited by B. L. Turner, E. Frossard, and D. S. Baldwin, pp. 113–132, CABI Publ., Wallingford, UK.

Celi, L., G. De Luca, and E. Barberis (2003), Effects of interaction of organic and inorganic P with ferrihydrite and kaolinite-iron oxide systems on iron release, Soil Sci., 168(7), 479–488, doi:10.1097/01.ss.0000080333.10341.a4.

Clark, L. L., E. D. Ingall, and R. Benner (1998), Marine phosphorus is selectively remineralized, Nature, 393(6684), 426–426, doi:10.1038/30881.

Clark, L. L., E. D. Ingall, and R. Benner (1999), Marine organic phosphorus cycling; novel insights from nuclear magnetic resonance, Am. J. Sci., 299(7-9), 724– 737, doi:10.2475/ajs.299.7-9.724.

Defforey, D., B. J. Cade-Menun, and A. Paytan (2016), A new solution 31P NMR sample preparation scheme for marine sediments, Limnol. Oceanogr. Methods, (in review).

Diaz, J., E. Ingall, C. Benitez-Nelson, D. Paterson, M. D. de Jonge, I. McNulty, and J. A. Brandes (2008), Marine polyphosphate: a key player in geologic

110 phosphorus sequestration, Science, 320(5876), 652–655, doi:10.1126/science.1151751.

Expedition 336 Scientists (2012a), Initiation of long-term coupled microbiological, geochemical, and hydrological experimentation within the seafloor at North Pond, western flank of the Mid-Atlantic Ridge., IODP Proc., 336, doi:10.2204/iodp.pr.336.2012.

Expedition 336 Scientists (2012b), Methods, Proc. Integr. Ocean Drill. Program, 336, doi:10.2204/iodp.proc.336.102.2012.

Expedition 336 Scientists (2012c), Sediment and basement contact coring, Proc. Integr. Ocean Drill. Program, 336, doi:10.2204/iodp.proc.336.106.2012.

Expedition 336 Scientists (2012d), Site U1382, Proc. Integr. Ocean Drill. Program, 336, doi:10.2204/iodp.proc.336.104.2012.

Goldberg, S., and G. Sposito (1985), On the mechanism of specific phosphate adsorption by hydroxylated mineral surfaces: A review, Commun. Soil Sci. Plant Anal., 16(8), 801–821, doi:10.1080/00103628509367646.

Greaves, M. P., and M. J. Wilson (1969), The adsorption of nucleic acids by montmorillonite, Soil Biol. Biochem., 1(4), 317–323, doi:10.1016/0038- 0717(69)90014-5.

Greaves, M. P., and M. J. Wilson (1970), The degradation of nucleic acids and montmorillonite-nucleic-acid complexes by soil microorganisms, Soil Biol. Biochem., 2(4), 257–268, doi:10.1016/0038-0717(70)90032-5.

Heath, R. T. (2005), Microbial turnover of organic phosphorus in aquatic systems, in Organic phosphorus in the environment, edited by B. L. Turner, E. Frossard, and D. S. Baldwin, pp. 185–203, CABI Publ., Wallingford, UK.

Heinonen, J. (2001), Biological role of inorganic pyrophosphate, Kluwer Academic Publishers, Norwell, MA.

Inagaki, F. et al. (2015), Exploring deep microbial life in coal-bearing sediment down to ~2.5 km below the ocean floor, Science, 349(6246), 420–424, doi:10.1126/science.aaa6882.

Ingall, E. D., P. A. Schroeder, and R. A. Berner (1990), The nature of organic phosphorus in marine sediments: new insights from 31P NMR, Geochim. Cosmochim. Acta, 54(9), 2617–2620, doi:10.1016/0016-7037(90)90248-J.

111 Jones, D. S., B. E. Flood, and J. V. Bailey (2016), Metatranscriptomic insights into polyphosphate metabolism in marine sediments, ISME J., 10(4), 1015–1019, doi:10.1038/ismej.2015.169.

Kearse, M. et al. (2012), Geneious Basic: An integrated and extendable desktop software platform for the organization and analysis of sequence data, Bioinformatics, 28(12), 1647–1649, doi:10.1093/bioinformatics/bts199.

Kolowith, L. C., E. D. Ingall, and R. Benner (2001), Composition and cycling of marine organic phosphorus, Limnol. Oceanogr., 46(2), 309–320, doi:10.4319/lo.2001.46.2.0309.

Laarkamp, K. L. (2000), Organic phosphorus in marine sediments: chemical structure, diagenetic alteration, and mechanisms of preservation, PhD Thesis, MIT-WHOI Joint Program.

Ludwig, W. et al. (2004), ARB: a software environment for sequence data, Nucleic Acids Res., 32(4), 1363–1371, doi:10.1093/nar/gkh293.

Martínez, A., M. S. Osburne, A. K. Sharma, E. F. DeLong, and S. W. Chisholm (2012), Phosphite utilization by the marine picocyanobacterium Prochlorococcus MIT9301: Phosphite utilization by Prochlorococcus MIT9301, Environ. Microbiol., 14(6), 1363–1377, doi:10.1111/j.1462- 2920.2011.02612.x.

McDowell, R. W., I. Stewart, and B. J. Cade-Menun (2006), An examination of spin– lattice relaxation times for analysis of soil and manure extracts by liquid state phosphorus-31 nuclear magnetic resonance spectroscopy, J. Environ. Qual., 35(1), 293, doi:10.2134/jeq2005.0285.

Mewes, K., J. M. Mogollón, A. Picard, C. Rühlemann, A. Eisenhauer, T. Kuhn, W. Ziebis, and S. Kasten (2016), Diffusive transfer of oxygen from seamount basaltic crust into overlying sediments: An example from the Clarion– Clipperton Fracture Zone, Earth Planet. Sci. Lett., 433, 215–225, doi:10.1016/j.epsl.2015.10.028.

Mills, H. J., B. K. Reese, and C. S. Peter (2012), Characterization of microbial population shifts during sample storage, Front. Microbiol., 3, 49, doi:10.3389/fmicb.2012.00049.

112 Van Mooy, B. A. S., A. Krupke, S. T. Dyhrman, H. F. Fredricks, K. R. Frischkorn, J. E. Ossolinski, D. J. Repeta, M. Rouco, J. D. Seewald, and S. P. Sylva (2015), Major role of planktonic phosphate reduction in the marine phosphorus redox cycle, Science, 348(6236), 783–785, doi:10.1126/science.aaa8181.

Price, M. N., P. S. Dehal, and A. P. Arkin (2010), FastTree 2 – Approximately Maximum-Likelihood Trees for Large Alignments, PLOS ONE, 5(3), e9490, doi:10.1371/journal.pone.0009490.

Pruesse, E., J. Peplies, and F. O. Glöckner (2012), SINA: Accurate high-throughput multiple sequence alignment of ribosomal RNA genes, Bioinformatics, 28(14), 1823–1829, doi:10.1093/bioinformatics/bts252.

Rao, N. N., M. R. Gómez-García, and A. Kornberg (2009), Inorganic Polyphosphate: Essential for Growth and Survival, Annu. Rev. Biochem., 78(1), 605–647, doi:10.1146/annurev.biochem.77.083007.093039.

Roussel, E. G., M.-A. C. Bonavita, J. Querellou, B. A. Cragg, G. Webster, D. Prieur, and R. J. Parkes (2008), Extending the Sub-Sea-Floor Biosphere, Science, 320(5879), 1046–1046, doi:10.1126/science.1154545.

Ruttenberg, K. C. (2014), The Global Phosphorus Cycle, in Treatise on Geochemistry, vol. 10, edited by H. Holland and K. Turekian, pp. 499–558, Elsevier.

Ryan, W. B. F. et al. (2009), Global Multi-Resolution Topography synthesis, Geochem. Geophys. Geosystems, 10(3), Q03014, doi:10.1029/2008GC002332.

Sannigrahi, P., and E. Ingall (2005), Polyphosphates as a source of enhanced P fluxes in marine sediments overlain by anoxic waters: evidence from P-31 NMR, Geochem Trans, 6(3), 52–59, doi:10.1063/1.1946447.

Sannigrahi, P., E. D. Ingall, and R. Benner (2006), Nature and dynamics of phosphorus-containing components of marine dissolved and particulate organic matter, Geochim. Cosmochim. Acta, 70(23), 5868–5882, doi:10.1016/j.gca.2006.08.037.

Shang, C., D. E. Caldwell, J. W. B. Stewart, H. Tiessen, and P. M. Huang (1996), Bioavailability of organic and inorganic phosphates adsorbed on short-range

113 ordered aluminum precipitate, Microb. Ecol., 31(1), 29–39, doi:10.1007/BF00175073.

Suzumura, M., and A. Kamatani (1995), Mineralization of inositol hexaphosphate in aerobic and anaerobic marine sediments: implications for the phosphorus cycle, Geochim. Cosmochim. Acta, 59(5), 1021–1026, doi:10.1016/0016- 7037(95)00006-2.

Turner, B. L., M. J. Papházy, P. M. Haygarth, and I. D. Mckelvie (2002), Inositol phosphates in the environment, Philos. Trans. R. Soc. B Biol. Sci., 357(1420), 449–469, doi:10.1098/rstb.2001.0837.

Turner, B. L., B. J. Cade-Menun, L. M. Condron, and S. Newman (2005), Extraction of soil organic phosphorus, Talanta, 66(2), 294–306, doi:10.1016/j.talanta.2004.11.012.

Turner, B. L., A. W. Cheesman, H. Y. Godage, A. M. Riley, and B. V. L. Potter (2012), Determination of neo- and d-chiro-Inositol Hexakisphosphate in Soils by Solution 31P NMR Spectroscopy, Environ. Sci. Technol., 46(9), 4994– 5002, doi:10.1021/es204446z.

Vershinina, O. A., and L. V. Znamenskaya (2002), The Pho regulons of bacteria, Microbiology, 71(5), 497–511, doi:10.1023/A:1020547616096.

Wanner, B. L. (1993), Gene regulation by phosphate in enteric bacteria, J. Cell. Biochem., 51(1), 47–54, doi:10.1002/jcb.240510110.

White, R. H., and S. L. Miller (1976), Inositol Isomers: Occurrence in Marine Sediments, Science, 193(4256), 885–886, doi:10.1126/science.193.4256.885.

Zhang, Y. S., W. Werner, H. W. Scherer, and X. Sun (1994), Effect of organic manure on organic phosphorus fractions in two paddy soils, Biol. Fertil. Soils, 17(1), 64–68, doi:10.1007/BF00418674.

114 Figures and Tables

3100 3400 3800 4100 4500 3600 Depth (mbsl)

3800 52’ o 4200

48’ 22 4000 o

4400

U1382B 44’ N 44’ 22 o 22

0 1 2 km

o o o 46 10’ W 46 5’ 46 0’ Figure 3–1. Location of the North Pond field site, located beneath the North Atlantic Gyre. The color scales indicate water depth for each site in meters below sea level (mbsl). Contour line intervals are 100 m for North Pond and the location of Site U1382 is indicated with a white dote. Map created using the default Global Multi- Resolution Topography Synthesis basemap [Ryan et al., 2009] in GeoMapApp version 3.6.0 (http://www.geomapapp.org).

115

(a) U1382B 1H–3 (3.2 mbsf) (b) U1382B 7H–3 (56.7 mbsf)

OrthoP monoesters O – OrthoP diesters

– – O R – O – P – = O Orthophosphate – –

O – O – R – O – P – O – R – – O = P – – O O – Phosphonate Polyphosphate O – – O – O OH Pyrophosphate –

– – – – R – P – = O O = P – – O – P = – O O O

– – – O O – O O = P – – O – P – = O n O – O –

30 2010 0-10 -20 -3030 20 10 0-10 -20 -30 Chemical shift (ppm) Chemical shift (ppm)

Figure 3–2. Solution 31P NMR spectra of sample U1382B 1H-3 (3.2 mbsf, panel a), and U1382B 7H-5 (56.7 mbsf, panel b). Spectra are plotted with 7 Hz line broadening, and are plotted with the orthophosphate peak at the same height. The inset spectra (light grey) are plotted with 2 Hz line broadening and enlarged to better show details.

116

14 MRP )

-1 12 TP 10 mol g μ 8

P concentration ( 2

red red CFA CFA

U1382B 1H-3U1382B 7H-3U1382B 1H-3U1382B 7H-3 step 1 P step 1 P step 2 P step 2 P

Figure 3–3. Mean MRP and total P content in pretreatment extracts from steps 1 (Pred,

citrate–dithionite–bicarbonate and water extracts) and 2 (PCFA, sodium acetate and water extracts), along with standard deviations derived from replicate sediment sample splits. Total P concentrations were measured on an ICP-OES, while MRP concentrations were measured colorimetrically on a QuikChem 8000 automated ion analyzer.

117 Table 3–1. Genes related to P uptake and metabolism identified using metagenomic analyses of sample U1382B 7H-5 (59.1 mbsf)

Gene Gene name Gene ID Taxonomy (MEGAN/RefSeq)

scaffold_1262_1 Alphaproteobacteria; scaffold_1273_1 Caulobacter ppX Exopolyphosphatase Alphaproteobacteria; scaffold_15267_1 Sphingomonas Alphaproteobacteria; scaffold_1123_1 Bradyrhizobium Alphaproteobacteria; scaffold_1066_1 Caulobacter scaffold_18080_1 ppK Polyphosphate kinase Alphaproteobacteria; scaffold_8377_2 Rhizobiales scaffold_38142_1 scaffold_41747_1 Actinobacteria; Blastococcus scaffold_18394_1 Actinobacteria; Curtobacterium Alphaproteobacteria; scaffold_4902_1 Caulobacter Phosphite transport Alphaproteobacteria; ptxC scaffold_16663_1 system permease Bradyrhizobium Alphaproteobacteria; scaffold_29682_1 Methylobacterium scaffold_2135_1 scaffold_4610_1 Alphaproteobacteria; scaffold_1299_2 Bradyrhizobium NAD-dependent scaffold_6712_1 ptxD phosphite Alphaproteobacteria; dehydrogenase scaffold_11504_1 Caulobacter scaffold_6879_1 Unclassified scaffold_791_1 Alphaproteobacteria; scaffold_2842_1 Rhizobiales Phosphonoacetate Alphaproteobacteria; phnA scaffold_18589_2 hydrolase Sphingomonas scaffold_28814_1 Actinobacteria; Actinomycetales scaffold_880_3 Unclassified Phosphonate-binding Alphaproteobacteria; phnD scaffold_5397_1 periplasmic protein Caulobacter scaffold_15336_1 Alphaproteobacteria scaffold_8108_2 Alphaproteobacteria; phnJ C-P lyase scaffold_6460_1 Bradyrhizobium Alphaproteobacteria; scaffold_10739_1 Rhizobiales α-D-ribose 1- scaffold_30395_1 Alphaproteobacteria; methylphosphonate 5- scaffold_14787_1 Bradyrhizobium phnG triphosphate synthase Alphaproteobacteria; scaffold_5184_1 subunit Rhizobiales

118 scaffold_4332_1 scaffold_692_1 Alphaproteobacteria; scaffold_673_1 Bradyrhizobium scaffold_145_1 appA Acid phosphatase scaffold_618_1 Alphaproteobacteria; scaffold_6851_1 Rhizobiales scaffold_1237_1 Actinobacteria; Blastococcus Alphaproteobacteria; scaffold_3183_1 appB Alkaline phosphatase Bradyrhizobium scaffold_39062_1 Actinobacteria; Curtobacterium

scaffold_2482_1 scaffold_17604_1 Alphaproteobacteria scaffold_41329_1 Phosphate transport pstA system permease Alphaproteobacteria; scaffold_8780_1 Bradyrhizobium scaffold_3988_1 Alphaproteobacteria; scaffold_31142_1 Caulobacter scaffold_36431_1 Unclassified Phosphate transport pstB system ATP-binding scaffold_2537_1 Alphaproteobacteria protein Alphaproteobacteria; scaffold_132_3 Asticcacaulis scaffold_10470_1 scaffold_4818_1 scaffold_1778_2 scaffold_3636_1 Alphaproteobacteria; scaffold_10908_1 Bradyrhizobium scaffold_21095_1 scaffold_14114_1 scaffold_2586_1 scaffold_34622_1 scaffold_3462_1 Phosphate regulon scaffold_12940_1 phoB response regulator scaffold_694_1 scaffold_688_2 Alphaproteobacteria; scaffold_2184_2 Caulobacter scaffold_167_4 scaffold_4218_1 scaffold_142_2 scaffold_33397_1 scaffold_22594_1 Alphaproteobacteria; scaffold_9531_1 Rhizobiales scaffold_22304_1 scaffold_10058_1 Alphaproteobacteria; scaffold_29529_1 Rhodopseudomonas

119 Alphaproteobacteria; scaffold_14822_1 Sphingomonas Betaproteobacteria; scaffold_1934_2 Burkholderiales scaffold_5127_1 Actinobacteria; Blastococcus scaffold_2427_2 Actinobacteria; scaffold_21832_1 Geodermatophilaceae scaffold_17979_1 Actinobacteria; Modestobacter scaffold_3982_1 scaffold_28298_1 Bacterodoites scaffold_115_3 scaffold_82_1 Unclassified scaffold_176_1 ABC transporter scaffold_14158_1 Alphaproteobacteria; phoD periplasmic solute- scaffold_3831_1 Bradyrhizobium binding protein scaffold_1777_2 Alphaproteobacteria scaffold_1598_1 scaffold_1521_1 scaffold_1253_1 scaffold_6727_1 scaffold_3196_1 scaffold_752_1 scaffold_8236_1 Alphaproteobacteria; scaffold_4017_1 Bradyrhizobium scaffold_2480_1 scaffold_2313_1 scaffold_3025_1 scaffold_4620_1 scaffold_3106_1 scaffold_10067_1 scaffold_633_2 scaffold_1080_1 Phosphate regulon phoR scaffold_351_2 sensor histidine kinase scaffold_784_1 scaffold_435_1 scaffold_2154_1 Alphaproteobacteria; scaffold_2015_1 Caulobacter scaffold_12374_1 scaffold_3364_1 scaffold_532_1 scaffold_176_2 scaffold_1162_2 scaffold_23_3 scaffold_142_3 scaffold_1720_1 Alphaproteobacteria; scaffold_3940_1 Phenylobacterium Alphaproteobacteria; scaffold_21167_1 Sphingomonas Betaproteobacteria; scaffold_27239_1 Burkholderia

120 scaffold_3982_2 Actinobacteria; Modestobacter scaffold_28543_1 scaffold_13755_1 Unclassified scaffold_7797_1 scaffold_2297_2 Alphaproteobacteria; Belnapia scaffold_1899_2 Alphaproteobacteria; Bosea scaffold_20108_1 scaffold_5595_1 scaffold_4327_1 scaffold_13144_1 Alphaproteobacteria; scaffold_1505_2 Bradyrhizobium scaffold_3404_1 scaffold_2390_1 sn-glycerol 3- scaffold_15028_1 phosphate transport scaffold_1604_1 ugpC Alphaproteobacteria; system ATP-binding scaffold_1274_2 protein Caulobacter scaffold_16604_1 Alphaproteobacteria; scaffold_8555_1 Rhizobiales scaffold_22893_1 scaffold_19336_1 Alphaproteobacteria; Rhizobium scaffold_8291_1 Actinobacteria; scaffold_14868_1 Geodermatophilaceae scaffold_13394_1 Actinobacteria; Modestobacter scaffold_14411_1 scaffold_11044_1 Unclassified

121 Table 3–2. Phosphorus compounds identified from 31P NMR spectral analyses of samples U1382B 1H-3 (3.2 mbsf) and 7H-5 (56.7 mbsf). The percentage values reported are relative to total sample P, including P removed during the 2–step pretreatment and residual P.

U1382B 1H-3 U1382B 7H-5 -1 % of sample TP -1 % of sample TP (nmol g ) (nmol g ) Phosphonates 6.71 0.05 2.19 0.02 Orthophosphate 143 0.99 64.7 0.51 Monoesters

α-glyc 1.53 0.01 – – β-glyc 3.05 0.02 – – scyllo IHP 1.53 0.01 – – P choline 1.53 0.01 – – Phytate 4.58 0.03 – – Neo IHP 3.05 0.02 – – Chiro IHP1 4.58 0.03 – – Chiro IHP2 4.58 0.03 – – Nucleotides 3.05 0.02 – – gluc 6 PO4 3.05 0.02 – – Mono1 3.05 0.02 1.80 0.01 Mono2 15.0 0.10 6.32 0.05 Mono3 3.05 0.02 1.29 0.01 Diesters

DNA 3.05 0.02 – – Other diester 6.10 0.04 2.19 0.02 Pyrophosphate 76.9 0.54 42.9 0.34 Polyphosphates 18.0 0.13 7.48 0.06

122 Table 3–3. Extraction yields for each step. Pred refers to P solubilized during the reductive step (step 1), PCFA refers to P released during the sodium acetate step (step 2). Porewater MRP concentrations are from Expedition 336 Scientists [2012c]. NMR extract TP corresponds to the total P content in the alkaline extracts (step 3). Extracted P is the fraction of P that was extracted in steps 1-3.

Average Porewater NMR Residual Sample P P Extracted depth MRP red CFA extract TP P (µmol ID (µmol/g) (µmol/g) P (%) (mbsf) (µM)a (µmol g-1) g-1) U1382B 3.2 1.79 9.82 3.88 0.31 0.36 97.5 1H–3 U1382B 56.7 1.35 8.87 3.49 0.13 0.17 98.7 7H–5 a Expedition 336 scientists [2012]

123 Table 3–4. Aluminum, calcium, iron, potassium, magnesium, manganese and 31 phosphorus content in P NMR extracts. T1 values are derived from P to iron and manganese ratios (w/v) using the relationship derived by McDowell et al. [2006].

Depth Al Ca Fe K Mg Mn P Calculated Sample ID P/(Fe+Mn) (mbsf) ————————µmol g-1———————— T1 (s) U1382B 1H–3 3.2 1.16 8.86 0.03 0.03 1.05 0.02 0.31 3.01 1.17 U1382B 7H–5 56.7 0.68 8.86 0.02 0.02 0.33 0.01 0.13 2.00 0.89

124 Figure S3–1. 16S rRNA phylogenetic tree for sample U1382B 7H-5 (56.7 mbsf) constructed from the metagenome.

Rhodopseudomonas palustris str ATCC 17001 Alphaproteobacteria Phreatobacter oligotrophus str PI_21

Rhodopseudomonas faecalis str gc Bradyrhizobium neotropicale str BR 10247

Nitrobacter hamburgensis str X14

Bradyrhizobium lablabi str CCBAU 23086 KC682406 Methylobacterium radiotolerans str DSM 1819

JN717169

Bradyrhizobium valentinumAfipia strmassiliensis LmjM3 str 34633

Bradyrhizobium elkanii str NBRC 14791 EF648045

F007948 KC682149 KX447592 KP182152

GQ103050 608

AB974254 KF003165

KC682133

FJ753573 Methylobacterium gregans str 002-074 Methylobacterium radiotolerans str 0-1 2125

1012 GQ417446 FJ787328

HQ853453 GU552885 Methylobacterium longum str 440 DQ310802

Rhizobium qilianshanense str CCNWQLS01 DQ310798 KC236648 DQ310800 U89820 JN648911 KX022837 EU488751 JQ660255 CP000133 Rhizobium etli str NBRC 15573 Methylobacterium aerolatum str 5413S-11KX443719 U28916 D14513 Nitratireductor basaltis str J3 2270 Aquamicrobium lusatiense str S1 Aquamicrobium defluvii str DSM 11603 SphingomonasSphingomonas oligophenolica glacialis strainKX507670 str C16y S213 Blastococcus saxobsidens str BC448 JN896610 1727 Phyllobacterium loti str S658 Geodermatophilus obscurus str DSM 43160 Sphingomonas echinoides str NBRC15742 Phyllobacterium bourgognense str STM 201 EU928863 276 EU170548 FJ716017 Blastococcus aggregatus str DSM 4725 FJ214361 LN555115 Sphingomonas echinoides st DSM1805 HM366501 JX438918 KM507603 KU519613 KC554955 AB696235 LC055582 Propionibacterium avidum str DSM 4901 KJ998636 Propionibacterium acnes str ATCC 6919 2288 KX131215

KP967484 HQ806563 D85995 GU269546 473 KC122613 0.04 HF968439 JF124186 EF639383 FJ195989 Actinobacteria JQ799959 Propionibacterium jensenii str DSM 20535 KC682173 KC682132326 HG315674 Propionibacterium granulosum str DSM 20700 Clavibacter michiganensis str LMG 7333 JN874334 Clavibacter michiganensis subsp. phaseoli str LPPA 982 AY375122447 EU861941 EU491091 GQ261814 AF348975 AB059259 HQ396620 AB094412 KJ184990 HQ143312 AY167857 Colwellia piezophila str Y223G JN018464 HQ225163 EU286985 GU269547 EU286985 FR872421 GU061318 EU977722 69 Colwellia psychrotropica str ACAM 179 FJ976536 ColwelliaColwellia polaris arctica str 539=7 str 435 JQ977207 Colwellia aquaemaris str S1 Enterobacter asburiae str JCM6051

AF025960KM979040 Leifsonia poae str VKM Ac-1401 Colwellia meonggei str MA1-3 AB428928 JX170711 Enterobacter cloacae str 279-56 1510 KC122712 KP267834 AY373406 JN398669KC768785

Enterobacter aerogenes str NBRC

AB741688 JN625723 414 KJ482890 KM880162 StaphylococcusStaphylococcus succinus vitulinus subsp. str casei ATCC str 51145SB72 Colwellia psychrerythraeaVibrio str ATCC panuliri 27364 str LBS2 Staphylococcus haemolyticus str JCM 2416 Staphylococcus jettensis str SEQ110 AB845252

AB920798 JQ427680 Staphylococcus devriesei str KS-SP_60 AM697310 EU563349 KJ873966 JQ427616 Vibrio azureus str LC2-005 KF081333

Vibrio harveyi str NBRC 15634 1053 KF071838

Vibrio alginolyticus str NBRC 15630 EF446898

KC431803

Cedecea neteri str GTC1717

Enterobacter cancerogenus str LMG 2693 Desulfotomaculum hydrothermale Lam5

KF500027

KJ480297

Gammaproteobacteria Firmicutes

125 CONCLUSIONS

The nature of sedimentary P and the extent of biotic and abiotic alterations that occur after sediment deposition in the open ocean are key aspects of the global marine P cycle that have important consequences for deep sedimentary subseafloor microbial life and reactive P burial efficiency. This thesis explores sedimentary P biogeochemical cycling by using an interdisciplinary approach to characterize the nature of sedimentary inorganic and organic P, and the numerous diagenetic processes affecting reactive P burial and sedimentary microbial metabolic processes.

Chapter 1 presents a new method for preparing sediment samples for solution 31P

NMR that enhances the use of this tool with organic matter–depleted sediments by amplifying the signal of low abundance P forms, including organic and biogenic P compounds. This is achieved through a two–step pretreatment prior to the solution 31P

NMR extraction that quantitatively removes orthophosphate, thus increasing the signal of low abundance P forms relative to orthophosphate. High-resolution P geochemistry records from two well-studied on the flanks of the Juan de Fuca Ridge and Mid-Atlantic Ridge are presented in Chapter 2, describing some of the key diagenetic processes altering solid-phase P at these sites. The deepest solution 31P

NMR spectra of marine sediments –to our knowledge– are presented in Chapter 3, along with metagenomic analyses. This innovative, interdisciplinary project reveals the diversity of organic P in oligotrophic open ocean sediments, as well as some potential microbial pathways for P uptake and metabolism in this environment.

126 Each of these studies constitutes a significant advancement over previous work and enhances our understanding of P cycling in the deep sedimentary subseafloor environment. In the first chapter, we propose a new sample preparation scheme that allows the use of an effective tool, solution 31P NMR, on samples with very low organic and biogenic P content. The majority of 31P NMR studies of marine sediments until now have focused on coastal sediments or sediments underlying high productivity regions of the world’s oceans, with higher organic to inorganic P ratios.

This new method is effective on oligotrophic sediments, and allows the detection of a wide variety of compounds, including orthophosphate monoesters, diesters and pyrophosphate. This new sample preparation scheme will contribute to the narrowing of the knowledge gap that exists on the nature and dynamics of organic and biogenic

P in open ocean sediments. The second chapter uses a well-established sequential extraction scheme for marine sediments (SEDEX) to characterize solid-phase P in ridge–flank systems at a high resolution. In addition to contributing to the field of marine P biogeochemistry, results from this chapter will be of value to the numerous microbiological investigations underway at those extensively studied sites. The third chapter utilizes the method for solution 31P NMR developed in chapter 1, and applies it to oligotrophic sediments in the North Atlantic, along with metagenomic analyses, to gain insights into the biotic processes altering organic P substrates in deep-sea sediments. We show how specific orthophosphate monoester compounds such as inositol hexakisphophate stereoisomers, glucose-6-phosphate, choline phosphate, and glycerophosphate decrease in relative abundance with depth. We also show evidence

127 of phosphonate, orthophosphate diester and polyphosphate remineralization with depth as a result of changing redox conditions, which may facilitate microbial degradation of these compounds. Lastly, we show putative microbial P uptake mechanisms in sediments that have been shown to host active microbial communities, supporting the hypothesis that some of the mechanisms presented may be utilized by active microorganisms.

Some possible new directions for studies of sedimentary P cycling in the deep subseafloor environment include the quantification of expressed P functional genes using quantitative reverse transcription polymerase chain reaction with specific primers designed using information from metagenomic analyses, and comparisons of expressed gene abundances with P geochemical data such as 31P NMR analyses and stable oxygen isotopes of phosphate. Alternatively, metatranscriptomic analyses, combined with enzymatic assays of deep–sea sediments could also provide valuable insights into the active enzymatic pathways influencing sedimentary P. The application of the sample preparation scheme for 31P NMR presented in chapter 1 at a high-resolution downcore would provide insightful information on the small-scale variations of sedimentary P forms (both inorganic and organic), which could not be gleaned from two spectra alone. These projects have the potential to impact related work on organic P biogeochemistry in the environment and current efforts seeking to constrain the extent of the activity of the deep biosphere.

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