Reclaiming phosphate from waste solutions with Fe(III

Reclaiming phosphate from waste solutions with Fe(III) –. 2 polysaccharide hydrogel beads for photo-controlled-release fertilizer. 3. M. H. Jayan S...
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Agricultural and Environmental Chemistry

Reclaiming phosphate from waste solutions with Fe(III) – polysaccharide hydrogel beads for photo-controlled-release fertilizer M. H. Jayan Savinda Karunarathna, Zachery R Hatten, Kerri M Bailey, Evan T Lewis, Amanda L Morris, Autumn R Kolk, Jenna C Laib, Nathan Tembo, Richard A Williams, Benjamin T Phillips, Bethany L Ash, W. Robert Midden, and Alexis D Ostrowski J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.9b02860 • Publication Date (Web): 15 Aug 2019 Downloaded from on August 17, 2019

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Reclaiming phosphate from waste solutions with Fe(III) – polysaccharide hydrogel beads for photo-controlled-release fertilizer

3 4 5

M. H. Jayan S. Karunarathna, Zachery R. Hatten, ‡† Kerri M. Bailey, ‡† Evan T. Lewis,† Amanda L. Morris,† Autumn R. Kolk,† Jenna C. Laib,† Nathan Tembo,† Richard A. Williams III, † Benjamin T. Phillips,† Bethany L. Ash, W. Robert Midden, Alexis D. Ostrowski*


Department of Chemistry and Center for Photochemical Sciences, Bowling Green State University, Bowling Green, Ohio


43403, USA


8 9

ABSTRACT: Photo-responsive hydrogels from polysaccharides and Fe(III) were used as a new system to capture and release


PO43- from waste solutions. Uptake of 0.6-1.5 mg phosphate per gram of hydrogels was determined from 800 ppm phosphate


solutions (pH 4.0-11.0). These beads also captured 1.2 mgg-1 phosophate from animal waste (raw manure, 727 ppm


phosphate, pH 7.6) which accounted for above 80% phosphate uptake. Irradiation of phosphate-loaded hydrogels degraded


the gels due to the photochemistry of the Fe(III)-carboxylates, giving controlled phosphate release (~81% after 7 days). No


release (< 2% after 7 days) was seen in the dark. Kale plant trials showed complete degradation of the hydrogels in ~2 weeks


under greenhouse conditions. Biomass analysis of kale treated with phosphate-loaded beads compared to controls indicated


no signs of toxicity. These results show Fe(III)-polysaccharide hydrogels were able to reclaim phosphates from waste


solutions and can be used as a controlled release fertilizer.

18 19 20

KEYWORDS: Controlled-release fertilizer, Fe(III) photochemistry, Photo-responsive hydrogels, Photodegradation, Phosphate


recycling, Green chemistry

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Fertilizers are a major component in any plan to achieve maximum crop yields and are considered a requirement for the agriculture


industry. Over the last five decades, the global fertilizer consumption has skyrocketed several magnitudes with the expansion of


agriculture.1–3 Phosphate is one of the important macronutrients for plants and crops and it is ubiquitous in soil.4 Currently phosphates


are mined at a global rate of around 20 million metric tonnes per year and almost all the phosphorus used in agriculture is from these


mined phosphates.5 However, it is speculated that the mined rock phosphates will be completely depleted in the next 50-100 years.6


Due to these phosphates being a limited resource, it is important to recycle and use phosphates efficiently. One way of recycling


phosphate is reclamation of phosphate from animal waste and using them in agricultural fields as fertilizers.7–10 Reclaiming and


recycling phosphate from animal waste is an attractive solution, because agriculture and livestock are major sources of phosphate


pollution. Globally, the production of primary and manufactured goods and services from crops and livestock accounts for 91% of


the world’s total blue water consumption (including fresh surface and ground water).11 In addition, when animal waste is used as


fertilizer, not all of the phosphate and nitrate nutrients are absorbed by the plants. Depending on soil type, rainfall, and elevation,


significant amounts of phosphate can leach into the environment, contributing to an array of ecological and human health problems


such as harmful algal blooms.12,13 One way of addressing these environmental issues associated with phosphate runoff is through the


use of controlled release fertilizer systems. Many of these controlled release fertilizer systems use synthetic polymers in combination


with natural substances which can slow waste and pollution rates.14–22 The use of petroleum-based materials in fertilizer systems and


application in agricultural fields could result in some other environmental issues. Therefore, another possible solution to these issues


is recycling nutrients such as phosphates from wastewater and animal waste generated from agricultural fields, animal farms and


certain industries using natural materials. These recycled nutrients can then be applied to agricultural fields as fertilizer.

41 42

Figure 1. A) Photochemical reaction between Fe(III) and Polyuronate B) Photo controlled phosphate release from Fe(III)-alginate hydrogel




Several groups have addressed the negative environmental impact of phosphate pollution by recycling phosphates with different


approaches. These methods include phosphate removal from wastewater by electrodialysis,23 electrocoagulation,24 hybrid


microfiltration-forwards osmosis membrane bioreactor process25 and using different absorbent systems developed with zeolites,14,26


red mud27 and hydrogels.28 In addition, methods for phosphate recovery from animal manure have also been developed.7,10

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Unfortunately, there is not controlled release on demand from these systems despite that many of these methods function well at


capturing phosphate. We have previously shown controlled delivery of small molecules from hydrogels using Fe(III)-carboxylate


photochemistry in polysaccharide-based hydrogel materials.29 The photochemical reaction between Fe(III) and polyuronic acids


showed controllable degradation and release. We used this as an inspiration to develop Fe(III)-polysaccharide gels that could absorb


phosphate from waste and show controlled release with light (Figure 1). By using biodegradable and relatively low-cost


polysaccharides, we can create controlled photo-release fertilizers that could be a potential solution to the environmental issues that


stem from animal agriculture. Herein, we use different polysaccharide-based hydrogel beads for phosphate uptake from artificial


aqueous waste solutions and raw animal waste solutions.


hydrogel beads at different pH values were studied. Kale plant trials were carried out with the phosphate loaded hydrogels to


demonstrate the light-controlled phosphate release under natural conditions and the biodegradability of the fertilizer system under


greenhouse environments.

The phosphate uptake capacity of different types of polysaccharide

59 60



Materials: Sodium alginate 35% mannuronate, Mw = 97,000 Da (Alginate G) and sodium alginate 61% mannuronate Mw =


110,000 Da (Alginate M) were received with thanks from Kimica Corporation. Alum (powdered, Kroger brand) was purchased from


a grocery store. Sodium phosphate (Na3PO4) anhydrous, disodium hydrogen phosphate (Na2HPO4) anhydrous, sodium dihydrogen


phosphate (NaH2PO4) anhydrous were purchased from Fischer scientific. Chitosan Mw 50,000 – 190,000 Da (Lot STBH6262) was


purchased from Aldrich chemical company. Pectin from citrus peel with 74% galacturonic acid (Lot SLBN9007V), Fe(III) chloride


hexahydrate > 98%, hydroxylamine hydro chloride 99%, 1,10 phenanthroline > 99% and sodium molybdate > 98% were purchased


from Sigma Aldrich. L-ascorbic acid > 99% was purchased from J. T. Baker chemical company. Any other chemicals used were


analytical reagent grade from Sigma-Aldrich or Fischer Scientific and all the aqueous solutions were prepared with de-ionized water.


Raw manure solutions were obtained from a concentrated animal feeding operation for dairy cattle in Putnam County, Ohio. These


raw manure solutions were mostly solution phase, with ~3.3% as suspended solids. The manure solutions used for these experiments


had an average pH of 7.6 ± 0.1 and average phosphate, ammonium and nitrate concentrations of 727 ppm, 1417 ppm and < 13 ppm


(less than minimum detection limit) respectively and were used as received without any additional treatment. Kale seeds (Dwarf blue


curled vates) from Burpee (W. Atlee Burpee and Co.) and the Sungrow professional growing mix (Fafard 4 Mix Metro Mix 510)


were used for the plant trials.


Hydrogel bead preparation: Gel bead preparation was done according to a method previously described with slight modifications


accordingly.29 In brief, 1% by weight aqueous polysaccharide solutions were used for hydrogel bead preparation. For the gel beads


with two polysaccharides, the solutions were prepared with equal weights from each polysaccharide so that total weight of


polysaccharides in solution is still 1%. For the preparation of the alum beads, first alum was dissolved in water to obtain a 0.1% by


weight alum solution. Then alginate 35% mannuronate (1% by solution weight) was slowly dissolved in the alum solution on the

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vortex to obtain the alum-alginate solution. These polysaccharide solutions were loaded into syringes with a needle of 20 gauge


except for chitosan solutions where 18 gauge needles were used due to the larger particle size of chitosan. Hydrogel beads were


obtained by dropping the polysaccharide solution into a petri dish filled with the FeCl3 solution of desired concentration (0.1 M, 0.05


M or 0.01 M) from a height of about 6 inches. Gel beads were allowed to sit in the FeCl3 solution for about 5 minutes for Fe(III)


coordination and then filtered. All the gel beads were rinsed with de-ionized water and allowed to air dry for 1 hour at 25˚C on top


of paper towels (10-20% relative humidity) before weighing the required amounts for the experiments.


Scanning Electron Microscopy (SEM): Hydrogel beads were cut into halves soon after freezing with liquid nitrogen and


lyophilized using a Labconco Freeze dry system / Freezone 4.5 machine. Dried samples were sputter coated with Au/Pd using


Hummer VI-A Sputter Coater for 2.5 minutes. SEM images were collected on a Hitachi S2700 scanning electron microscope at 12


kV. Energy Dispersive X-ray Spectroscopy (EDS) was performed using EDAX detecting unit (model: PV77-47700-ME) at a voltage


of 20 kV.


Hydrogel composition analysis: For water content analysis, an exactly weighed sample of hydrogel beads was allowed to dry in


an oven at 45˚C until constant weight was achieved. The final weight after the drying process was recorded and the water content


was calculated based on weight loss on drying. For iron content analysis, exactly weighed 10.0 g samples of gel beads were photolyzed


using a Thorlabs 405 nm light emitting diode (LED) source with a power of 50 mWcm-2 and any gel bead particles that did not


degrade completely after 3 days of irradiation were chopped with a mortar. The irradiated solution was transferred into a 100 mL


volumetric flask and 1 mL of 10% hydroxylamine hydrochloride was added to convert any remaining Fe(III) to Fe(II) and diluted up


to the mark. Quantification of Fe(II) was done by colorimetric method where 0.1 mL of the diluted solution was added into a 10 mL


volumetric flask which contained 2 mL of pH 4.0 acetate buffer (prepared by diluting 600 mL of 1 M sodium acetate and 360 mL of


0.5 M H2SO4 to 1 L and adjusting the pH). Then a 1 mL aliquot of 0.1% by weight 1,10 phenanthroline solution was added and


diluted up to mark. The solution was allowed to stand for 15 minutes for color development and absorbance of the colored species


was recorded at 510 nm corresponding to Fe(II) phenanthroline complex.

102 103

FTIR Spectroscopy: Freeze dried hydrogel samples were used for FTIR spectroscopy and a Jasco FTIR-4000 machine equipped with a single reflection ATR accessory was used for spectrum collection.


Phosphate uptake from artificial waste solutions: Hydrogel bead samples of 10.0 g were placed in 50 mL glass beakers with 20


mL solutions of desired phosphate solution (Na3PO4, Na2HPO4, NaH2PO4 or pH = 7 phosphate buffer) with the phosphate


concentration of either 800 ppm or 100 ppm PO43-. Beakers were covered with parafilm and placed on top of Daigger 22407A


mechanical shaker under dark conditions for 24 hours. Then the gel beads were filtered off and the phosphate solutions were used for


remaining phosphate analysis.


For the experiments with 800 ppm phosphate solutions, the filtrates after the soak process were diluted 100 times so that the final


phosphate concentration of the solutions was below 6 ppm PO43-. Colorimetric analysis was performed using a method designed by


modifying the US Environmental Protection Agency (EPA) test method for orthophosphates using UV-Vis spectroscopy.i In the

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current method a molybdate reagent was prepared by dissolving 1 g (4.86 mmol) of sodium molybdate in 40 mL of 3 M sulfuric acid


and 0.2 g of ascorbic acid was added to the solution. A 3 mL portion of this molybdate reagent was added into each test tube containing


20 mL of diluted phosphate solutions and they were covered with parafilm. The test tubes were placed in a hot water bath at 60˚C for


30 minutes and allowed to cool to room temperature before measuring their absorbance of the phosphomolybdate complex at 830


nm. Remaining phosphate concentrations were calculated using a calibration plot prepared for absorbance at 830 nm for phosphate


solutions with 0.0 – 6.0 ppm phosphate concentration range (Figure S1). Phosphate uptake by the gel beads was obtained by the


difference of initial and final phosphate concentration of the solution.


All the solutions for experiments with 100 ppm PO43- solutions were analyzed as it is without any dilutions since the remaining


concentration of phosphate is low enough for the instrument. Remaining phosphate content was analyzed with the SEAL Analytical


AQ2+ Discrete Chemical Analyzer (AQ2+) with a range of application from 0.005 – 1.0 mg P/L (0.015 – 3.0 ppm PO43-) using the


US EPA test method 365.1.ii


Phosphate uptake from manure solutions: Exactly weighed 10.0 g samples of gel beads were placed in 100 mL plastic cups and


20 mL of raw manure solutions were added to them. The cups were allowed to stand for 24 hours and then the manure solutions were


separated from the beads. Remaining phosphate content in the manure solution was analyzed using the SEAL Analytical AQ2 Discrete


Analyzer. Phosphate analysis of raw manure solution was also performed after a 50 X dilution to bring the concentration into the


working range of the instrument.


Light controlled phosphate release in water: Samples of 10.0 g from alginate M-0.1 M FeCl3 beads were soaked in 20 mL of 800


ppm phosphate solution (pH = 7.0) for 24 hours. These beads were rinsed with 20 mL of DI water immediately and placed in petri


dishes with 20 mL of water and covered with a glass petri dish. The first set of samples was stored under dark conditions at room


temperature and the second set of samples was placed on the roof to expose to sunlight (on November 2017 at 41.6°N latitude). The


samples exposed to sunlight experienced an average day time high temperature of 5.9 °C with mostly clear skies. The third set of


samples was irradiated with the Thorlabs 405 nm LED light with an irradiance power of 50 mWcm-2 (measured with a S121C -


Standard Photodiode Power Sensor, 400 - 1100 nm, 35.5 mW) and the laboratory temperature of 25 °C. Aliquots of 6 mL were


collected from each of these petri dishes every 24 hours for analysis and 6 mL of de-ionized water was added each time to maintain


the same volume. The experiment was continued for 7 days and the solutions were diluted appropriately and analyzed for phosphate


content using the same colorimetric molybdate method used for the phosphate uptake experiments. Fe(II) content of each of these


solutions was also analyzed using the same 1,10 phenanthroline method which was used in hydrogel composition analysis.


Plant trials: Kale seed germination studies were carried out under greenhouse conditions in small pots filled with 5 g of Sungrow


professional growing mix 510 and dwarf blue curled vate kale seeds (Brassica oleracea) were placed about 1 cm below the surface.


Three different conditions were provided for the seeds with 8 pots under each condition. As the first condition, each pot was treated


with 5.0 g of Alginate G – 0.1% alum hydrogels which were soaked in a 800 ppm phosphate solution (pH = 7) for 24 hours. Portions


of 5.0 g of hydrogel beads without soaking in phosphate solution were used as the second condition and the third condition was pots

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with no hydrogel beads added. All the pots were given equal amounts of de-ionized water three times every week to ensure adequate


water supply, and the experiment was carried out for three weeks.


To access the effects of the hydrogels on growth after germination, kale plant trials were also conducted under greenhouse


conditions. Kale plants which were 17 days old (from day of seed planting) were transplanted in plastic pots (8.5 cm x 10 cm diameter)


filled with 100 g of Sungrow professional growing mix 510. A thin layer of water-washed and dried sand (75 g) was added to hold


the gel beads on the top of the pot upon watering. Similar to the seed germination experiments, three conditions were provided for


the plants namely, 10.0 g of phosphate loaded Alginate 53% M – 0.1% alum beads, 10.0 g of control beads (no nutrient loading) and


control with no beads. 8 replicates under each condition were performed. Plants were watered three times every week with 100 mL


of de-ionized water. After three weeks, another set of gel beads was added for the gel bead conditions. By the end of the sixth week


from transplanting, kale plants were harvested and the root systems were separated from the plants. All the parts of the plants except


the root system were cut into pieces and weighed. Then they were oven dried at 45˚C until a constant weight was obtained and above


ground bio mass was determined.

156 157



To investigate the uptake of phosphate by Fe(III)-alginate hydrogels, beads were prepared with different types of alginates, and


with different concentrations of Fe(III). These beads are spherical and approximately 3-4 mm in diameter with an orange color


indicative of Fe(III) (Figures S2). Once prepared the Fe(III)-alginate beads were soaked in phosphate solutions of different


concentrations, and the amount of phosphate that remained in the beads was determined. Hydrogel beads were prepared under a


variety of different conditions to investigate the effects of Fe(III) concentration, the structure of the polysaccharide, and pH on the


overall uptake of phosphate into the gels.

Symmetric R-CO 2 symmetric R-CO 2 Stretching stretching 1451 cm-1

% Transmittance

100 O-H stretching broad band 3419 cm-1


O-H Stretching


asymmetric R-CO 2 2 Asymmetric R-CO -1 stretching 1618 cm Stretching Out-of-phase out –of-phase P-O Stretching P-O stretching


1417 cm-1


Alginate G-Alum beads Alginate G-Alum beads soaked in phosphate solution

75 4000


in-phase In-phase P-O stretching P-O1029 Stretching cm-1





Wavenumber / cm

1500 -1

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Figure 2. FTIR spectra of alginate – alum beads before and after phosphate uptake


FT-IR analysis of Alginate G-Alum-0.05 M FeCl3 beads (Figure 2) clearly showed characteristic peaks for O-H stretching (broad


band centered around 3400 cm-1), asymmetric stretching of R-CO2- (intense peak around 1600 cm-1), symmetric stretching of the R-


CO2- (peak around 1400 cm-1), and asymmetric C-O-C stretching (around 1100 cm-1). After soaking in phosphate solutions, clear


differences were seen at 1080 cm-1, 1650 cm-1 and 3300 cm-1 of the FTIR spectra (Figure 2). The most intense peak around 1080 cm-1


was attributed to the in-phase P-O stretching frequency of the PO43- group.30 Studies on FTIR spectra of phosphate ions have shown


that phosphates have a medium peak around 1650 cm-1 and a very strong, broad peak in the 3200 cm-1 region.31 An increased IR


absorbance was seen in these regions for gels, which were soaked in PO43-. Therefore, these observations in the FTIR spectra of gels


soaked in phosphate solution indicated that the phosphates ions were trapped inside the hydrogels during the soaking process.


In addition to visual observation of the hydrogel beads (Figure 3A), we studied the gel structure using electron microscopy.


Scanning Electron Microscope images of these gel beads showed that the surface of the gel beads had a crust-like structure with


mostly smooth areas (Figure 3B), by contrast the interior of the gel bead was more porous in nature (Figure 3C) with interconnected


channels to facilitate the large amount of water present in the hydrogel system. Such structure is typical of these kind of polysaccharide




Quantitative elemental analysis was performed for the hydrogel beads using Energy Dispersive X-ray Spectroscopy (EDS) before


and after soaking in the phosphate solution (Figure S3). It is important to note that the percent weight of each element was calculated


excluding the gold and palladium which were used to coat the samples before the experiment (Au and Pd appear at 2.12 and 2.84 in


EDS spectra respectively). EDS confirmed the presence of phosphorus in the hydrogel system after the soak time which was not seen


in the gel beads before soaking in phosphate (Table 1). Phosphorus was present more than 5% by weight in the gels after the soaking


process. Also upon the soaking process, oxygen became the most abundant element in the system due to the additional oxygen atoms


from PO43-.

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186 187

Figure 3. A) Optical image of alginate G - 0.1 M Fe beads, B) exterior and C) interior of Alginate G-Alum- Fe bead prepared with Alginate


53% M, alum and 0.05 M FeCl3 bead under SEM


Another interesting feature was that the EDS elemental analysis performed for the phosphate-soaked gel bead exterior showed


differences in the element percentages compared to the interior (Table S1). Higher carbon percentage in the gel exterior along with


higher iron, oxygen and phosphorus percentages in the gel interior could be due to the slight rearrangements of the dynamic hydrogel


structure during the soaking in phosphate solution. It can be assumed that to facilitate the negatively charged phosphate groups within


the gel system, positively charged Fe(III) ions tended to move towards interior of the dynamic hydrogel structure along with the


carboxylate groups that they were coordinated to. This exposed the carbon polysaccharide chain towards the gel surface which caused


the higher carbon content in the gel exterior compared to the interior. SEM images of hydrogels soaked in phosphate solution (Figure


S4) did not show any significant differences compared to the SEM images of hydrogel beads before soaking (Figure 3).


It was assumed that the phosphate uptake by the hydrogels would be largely controlled by the charge and Fe(III)-coordination in


the gels. To investigate the differences, phosphate uptake was determined from several kinds of polysaccharide gels with different


Fe(III) concentrations. All the different types of hydrogel beads showed phosphate uptake behavior of 1 mg per 1 g of gel beads or


more at pH 7 with the 800 ppm phosphate solution (Figure 4). Regardless of the polysaccharide composition, gel beads prepared with


0.1 M FeCl3 solution showed a phosphate uptake of 1.2 mg of phosphate per 1 g of gels or more at pH = 7 (Figure 4A). The slight


changes of phosphate uptake with the change of the polysaccharide composition was statistically insignificant at 95% confidence

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interval (p = 0.20, 0.12 and 0.14 for alginate G-0.1 M Fe(III) beads compared to alginate M-0.1 M Fe, alginate G-pectin-0.1 M Fe(III)


and alginate G-chitosan-0.1 M Fe(III) respectively). Also it was noted that the effect of the concentration of Fe(III) solution used for


the gel bead preparation towards the phosphate uptake behavior (Figure 4B) was also statistically insignificant at 95% confidence


interval for the three Fe(III) concentrations used (0.01, 0.05 and 0.1 M). All the gel beads prepared with the 0.1 M FeCl3 solution had


slightly higher phosphate uptake and stability over a wider pH range than the beads prepared with 0.01 M FeCl3 solution (Figure 4D


and S5). This could be because beads prepared with 0.1 M FeCl3 solution showed more Fe inside the beads than the beads prepared


with the more diluted solution (Table 2) suggesting that Fe(III) was definitely playing a role in phosphate uptake. It can be believed


that Fe(III) ions are involved in PO43- uptake in an iron-phosphate complex formation mechanism similar to the previously reported


solution phosphate uptake methods using other metal ions like magnesium.34


Table 1: Elemental analysis of Alginate G-Alum-0.05 M FeCl3 hydrogel bead interior before and after soaking in phosphate


solution Element

% weight before PO43-

% weight after PO43-


























In our previous work, we have shown that the Fe(III)-alginate beads have been shown to have nanoclusters with oxo, hydroxo, and


carboxylate-bridged di-nuclear Fe(III) moieties.35 These kinds of coordination moieties exist in nature such as active sites of some


phosphodiesterase enzymes, and this moiety has been shown to coordinate well to phosphate.36 Strong phosphate binding to Fe(III)


ions has also been observed in natural systems such as in purple acid phosphatases. Phosphates bind to the Fe(III) ion of the dinuclear


Fe(III) center of these enzymes with good affinity.37 Also iron is widely used in waste water treatment plants to recover phosphorus


as iron – phosphorus compounds. This is due to the many different mechanisms by which phosphorus can bind to iron including


formation of Fe-phosphate minerals and different phosphate ion binding mechanisms to iron oxides.4,38 We believe that the phosphates


are binding to the iron nanoclusters in the polyuronate-based hydrogel network system in a similar way to other iron(III) – phosphate


binding that occurs in nature.


It is known that phosphates can bind strongly to Fe(III) and aluminum oxy-hydroxides and these can play an important role in


nature as efficient carriers of phosphates.37,39 The increase in phosphate uptake in the alum-based beads compared to the Alginate G-


0.05 M Fe(III) beads can be attributed to the presence of Al(III) ions in addition to Fe(III) ions (Figure 4C).

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Figure 4. Phosphate uptake (mg of phosphate absorbed per g of gels) by alginate beads from 800 ppm phosphate solution changes depending


on A) type of polysaccharide B) concentration of FeCl3 solution used to prepare the hydrogels C) introduction of other cations (from alum)


in addition to the FeCl3 and D) change in pH of the phosphate solution. (pH = 7 for all other uptake studies)


In the case of Alginate G beads, the beads prepared with 0.05 M Fe(III) solution showed good phosphate uptake quite similar to the


beads prepared with 0.1 M Fe(III) (Figure 4B). Even the phosphate uptake from beads prepared with 0.01 M Fe(III) was not


significantly different from the uptake from 0.1 M Fe(III) beads at pH 7. When considering the Alginate G beads prepared with FeCl3


solutions with different concentrations, we saw that the total iron content was lowest for the beads prepared with 0.01 M Fe(III)


solution and the maximum iron content was seen for the beads prepared with the intermediate Fe(III) concentration of 0.05 M (Table


2). While the beads prepared with the 0.1 M FeCl3 solution had slightly less total iron content than in the beads prepared with the


0.05 M FeCl3 solution, this difference is not statistically significant at 95% confidence interval (p = 0.09). This follows the


observations that we saw in the phosphate uptake experiment where beads prepared with 0.1 M FeCl3 absorbing slightly less


phosphate than beads prepared with 0.05 M FeCl3 solution. Again this suggests a role for Fe(III) in the binding of phosphates within


the hydrogels.


These changes of total Fe content that we observed could be due to the presence of different Fe species within the hydrogel systems


prepared with different FeCl3 solutions in different ratios. In this study we didn’t focus on investigating the exact iron species and


only total iron content was studied to see whether it relates to phosphate uptake by different kinds of gel beads. In the future, we hope


to investigate more on the Fe(III)-phosphate species formed during this process.


Composition analysis was carried out for hydrogel beads to study their differences (Table 2). Water was the major constituent in


all the hydrogel beads accounting for more than 96% by weight for types of beads that were analyzed. Total Fe(III) content by weight


in the beads was 0.2% or less. The rest of the bead composition was the polysaccharide. It is interesting to see that hybrid Fe/alum

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beads showed higher Fe(III) content than analogous beads made with only FeCl3. This may be due to the presence of Al(III) ions


resulting in some mixed Fe(III)/Al(III) moieties inside the beads that contributed to a higher overall metal content.


The phosphate uptake process was mostly independent from the pH for almost all the gels prepared with high Fe(III) concentrations.


For the gels prepared with lower FeCl3, the phosphate uptake was decreased with increasing pH. (Figure 4D and Figure S5). In


addition, the Fe(III)-polysaccharide hydrogels showed some instability (especially the beads prepared with 0.01 M FeCl3 solutions)


after the soak time of the experiments with pH = 11.5 Na3PO4 solution. Some of the gel beads dissolved in the highly basic solution,


turning the solution orange color due to Fe(III) from the bead dissolution and dispersal into the bulk solution of Fe(III) species. This


dissolution of the gel beads at high pH resulted in less overall amount of gel beads for phosphate uptake, and this could account for


the relatively lower phosphate uptake values.


To access the effect of initial phosphate concentration, similar experiments were carried out with the lower concentration 100 ppm


phosphate solution (pH = 7), and they showed 99% of phosphate uptake from the solutions (Figure 5A). In the lower concentration


of phosphate solution, the hydrogel beads absorbed phosphates at about 0.2 mg of PO43- per 1 g of gel beads, which was about 1/5th


of the maximum phosphate uptake capability.


Once again the changes of phosphate uptake capability with the change of hydrogel type was not statistically significant at the 95%


confidence interval. The low phosphate concentration in solution did not allow the beads to uptake phosphate at their maximum


efficiency. But all types of gel beads tested were affected similarly regarding their binding of phosphate.

263 264 265 266

Table 2: Composition analysis of hydrogel beads as prepared Others

Gel bead type

% Water by

% Fe by

by difference

weight [a]

weight [b]


Alginate G-Alum96.17 ± 0.08

0.21 ± 0.01


97.09 ± 0.01

0.17 ± 0.08


98.1 ± 0.1

0.16 ± 0.02


97.16 ± 0.04

0.23 ± 0.04


0.05 M FeCl3 Alginate G-0.05 M FeCl3 Alginate M-0.1 M FeCl3 Alginate M-0.05 M FeCl3

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Alginate M-0.01 M 98.27 ± 0.05

0.04 ± 0.01



267 268

[a] % water determined by loss on drying at 45°C until constant weight is achieved, [b] % Fe determined by converting all the Fe(III) to Fe(II) and analyzing the Fe(II) content using 1,10 phenanthroline according to a previously reported method.40


Animal waste solutions were used as the next step to evaluate the phosphate uptake capability of these Fe(III)-polysaccharide beads


in this natural, multi-component system. Phosphate uptake experiments carried out with dairy liquid animal waste solution (with a


phosphate concentration of 727 ppm and a pH of 7.6 ± 0.1) also showed relatively good phosphate absorption behavior by the gels


(Figure 5 B). Similar to the phosphate uptake experiments with the 800 ppm phosphate solutions (pH = 7.0), the phosphate uptake


values were around 1.2 mgg-1.


Figure 5. Phosphate uptake by different hydrogel beads in A) 100 ppm phosphate solution (pH = 7.0) and B) Phosphate uptake from manure


solution with 727 ppm phosphate (average pH = 7.6 ± 0.1) compared to the phosphate uptake from 800 ppm phosphate solution (pH = 7.0)


The manure solution used for these experiments had an initial phosphate content of 727 ppm phosphate and a pH of 7.6 ± 0.1. The


hydrogel beads showed their maximum phosphate uptake behavior similar to the artificial phosphate waste solutions with 800 ppm


phosphate. Even though manure was a mixture of some other ions such as ammonium and nitrates and various suspended solids,


these did not significantly affect the phosphate uptake behavior of the gel beads.


The ability to uptake phosphate from solutions makes these hydrogels candidates for recycling phosphorus. The photochemical


reaction between Fe(III) and carboxylate groups breaks the polymer chain and eventually the hydrogel network, hence we could


release phosphate by using light. Phosphate loaded hydrogel beads showed very low phosphate release under dark conditions (Figure


6A and Table S2). After 7 days, the total phosphate release was less than 1.5% and the gel beads didn’t show any change in appearance


(Figure 6A and B). The samples exposed to sunlight showed a 15% phosphate release over 7 days, some of the hydrogel beads

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degraded and the remaining gel particles were dark gray due to the photoreaction. The samples irradiated with 405 nm light (50


mWcm-2) showed the highest phosphate release with greater than 80% of phosphate release after 7 days. Also, the remaining hydrogel


particles were less than the sunlight samples (Figure 6B). This significant difference in sunlight and 405 nm conditions was attributed


to lower intensity of light that would initiate the photoreaction.

289 290

Figure 6. A) Phosphate release from gel beads under different light conditions B) Gel bead degradation under different light conditions


The solar irradiation consisting of wavelengths from ultra-violet to infra-red was less intense than the narrow-band 405 nm LED


that we used in the lab. Additionally, the sunlight conditions were only irradiated for about 14 hours per day (during fall) whereas the


samples irradiated with 405 nm light source in the lab were exposed to the light for 24 hours per day. The temperature of the LED


experiment varied less in the lab than outdoors during day and night, and the average outdoor temperature was lower than the lab,


and that might also affect the phosphate release. The overall photochemical reaction yield was determined by quantifying the Fe(II)


generated during light irradiation. This trend in Fe(II) production followed the phosphate release trend (Figure S6), showing that as


we expected, the Fe(III)-carboxylate photochemical reaction degraded the hydrogel to generate Fe(II) and released the trapped


phosphate ions from the gels.


To investigate the application of these hydrogels as a solid fertilizer, plant trials on kale were performed in a green house. Seed


germination experiments were carried out with kale seeds to assess the toxicity of the Fe(III)-polysaccharide hydrogels for plants.


The kale showed no signs of negative influence from the gel beads towards their germination (Figure 7). It was observed that the gel


beads were stable under the greenhouse conditions for several days before degradation. Twenty days following gel bead application,


it was difficult to observe any gel beads in the pots because of dehydration and then decomposition of the beads, indicating the photo-


and chemo-degradability of our hydrogel fertilizer system.

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Hydrogel toxicity was further assessed during plant growth and maturation. Plant trials with kale showed promising similar growth


in the plants that were treated with the phosphate-loaded hydrogels compared to the untreated controls (Figure S7). Similar to the


germination studies, there were no visible signs of negative effects for the plants (discoloration, etc.) from our hydrogels (Figure S8).


During the first couple of weeks, kale plants treated with phosphate beads had an enhanced growth compared to the other plants as


observed visually. By the time of harvesting (6 weeks from transplanting), the plants treated with phosphate-loaded hydrogel beads


collectively showed only small differences in growth compared to the others.

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Figure 7. Kale seed germination experiment under three different conditions. Left: phosphate-loaded beads, middle: control beads, right:


just soil


Overall, comparison of the above root biomass of the plants not treated with any beads and the plants treated with hydrogels without


phosphates showed only small differences (p = 0.44 and 0.08 for biomass differences of control plants, plants treated with control


beads, and phosphate-loaded beads respectively) at the 95% confidence interval (Figure 8). The lack of difference could be due to


the presence of nutrients in the growing mix that we used such that phosphate was not limiting growth, or that the amount of phosphate


added to the plants treated with phosphate beads was insufficient for enhanced growth. Furthermore, plants require other nutrients


such as potassium and nitrogen sources other than phosphates, which we did not provide in this experiment. While enhanced growth


was not seen, our phosphate loaded hydrogel system has advantages over conventional fertilizer including non-toxic components,


ability to biodegrade, and controlled release mechanism using light. Future work involves studying the uptake of other inorganic ions


present in wastewater (ammonium, nitrate etc.) by these Fe(III)-polysaccharide hydrogel beads and their photo-controlled release


towards plants with much longer lifetime.

324 325

Figure 8. Above ground biomass analysis of kale plants grown under three different growth conditions

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In summary, we developed a novel system to capture phosphate ions from waste water and animal manure using natural


polysaccharide materials which can be used as a photocontrolled phosphate fertilizer for plants. Our Fe(III)-alginate hydrogel beads


consisted primarily of water with a small amount of Fe(III) coordinated inside. These beads showed good phosphate uptake in the pH


range of 4.0 – 9.0 within 24 hours in both model wastewater solutions and liquid animal waste. The uptake behavior slightly changed


with the polysaccharide composition and with the concentration of Fe(III) used for gel bead preparation. Exposure of these hydrogels


to light, triggered the Fe(III) – carboxylate photochemistry which resulted in degradation of the hydrogels and release of trapped


phosphates. Plant trials carried out with kale plants showed that these phosphate hydrogels were candidates as a light-controlled slow


release fertilizer system. This Fe(III)-polysaccharide hydrogel beads system is an environment-friendly approach for reclamation of


phosphate from wastewater and animal waste with an easily scalable process to create light-responsive solid fertilizers. Other than


the environmental benefits, farmers can utilize the waste and reduce their fertilizer cost with this easy method to generate their own


fertilizer on site.




Supporting Information


Supporting information including colorimetric phosphate calibration, SEM and EDS analysis, Fe(II) release, and Kale plant photographs,


and a table of phosphate release is available. This material is available free of charge via the Internet at




Corresponding Author


* E-mail: [email protected]


Author Contributions


The manuscript was written through contributions of all authors. / All authors have given approval to the final version of the manuscript.


†Undergraduate researchers, ‡ These authors contributed equally


Funding Sources


We acknowledge the Herman Frasch Foundation for Chemical Research in Agricultural Chemistry (811-HF17) for funding to support this


project. We acknowledge funding of this work by grant numbers 6833 and 7177 from the Research and Development program of the Ohio


Water Development Authority, grant number R/HAB-17-ODHE from the Harmful Algal Bloom Research Initiative of the Ohio


Department of Higher Education managed by Ohio Sea Grant, and grant number SG 538-2018 of the Ohio Lake Erie Commission.

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i US EPA test method accessed on 03/13/2018 from the link

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ii US EPA test method accessed on 03/13/2018 from the link


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We thank Dr. Helen J. Michaels, Department of Life Sciences, Bowling Green State University for helpful discussions on plant trials, and


we thank Frank A. Schemenauer, the BGSU horticulturist and greenhouse manager for help with planting and plant care instructions. We


thank Dr. Marilyn Cayer for help with Electron Microscopy. We thank the Kimica Corporation for kindly providing us with alginate. We


acknowledge the ACS Herman Frasch Foundation for funding for this project. We acknowledge funding of this work by grant numbers 6833


and 7177 from the Research and Development program of the Ohio Water Development Authority, grant number R/HAB-17-ODHE from


the Harmful Algal Bloom Research Initiative of the Ohio Department of Higher Education managed by Ohio Sea Grant, and grant number


SG 538-2018 of the Ohio Lake Erie Commission. ZRH acknowledges BGSU Department of Chemistry for a summer research fellowship.


KMB, ETL, ALM, ARK, JCL, NT, RAW, BTP acknowledge BGSU Center for Undergraduate Research for undergraduate research grants.




Alginate G, sodium alginate 35% mannuronate; Alginate M, sodium alginate 61% mannuronate; EDS, Energy Dispersive X-ray


Spectroscopy; EPA, Environmental Protection Agency; FT-IR, Fourier Transform Infra-Red; LED, Light Emitting Diode; SEM, Scanning


Electron Microscopy





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