Vol.3, No.6, 806-815 (2012) Agricultural Sciences
http://dx.doi.org/10.4236/as.2012.36098
Swine waste as a source of natural products:
A carotenoid antioxidant
Lawrence B. Cahoon1*, Christopher J. Halkides2, Bongkeun Song3, C. Michael Williams4,
George R. Dubay5, Alexandra Fries6, Johanna Farmer7, William Fridrich8,
Charles Brookshire9
1Department of Biology and Marine Biology, UNC Wilmington, Wilmington, USA; *Corresponding Author:
2Department of Chemistry and Biochemistry, UNC Wilmington, Wilmington, USA;
3Center for Marine Science, UNC Wilmington, Wilmington, USA;
4Animal and Poultry Waste Management Center, N.C. State University, Raleigh, USA;
5Department of Chemistry, French Family Science Center, Duke University, Durham, USA;
6Integration and Application Network, University of Maryland Center for Environmental Science, Cambridge, USA;
7Ecotoxicology Lab, BLD 218, Department of Biology, University of Louisiana at Lafayette, Lafayette, USA;
81918 Princess Street, Wilmington, USA;
9209 Stonewall Jackson Drive, Wilmington, USA;
Received 28 June 2012; revised 31 July 2012; accepted 9 August 2012
ABSTRACT
Development of Environmentally Superior Tech-
nologies for swine waste management has fo-
cused on extraction of products with relatively
low unit values. Analyses of the bacterial com-
position of swine waste lagoon samples con-
firmed the presence of several purple non-sulfur
bacteria (PNSB) species known to produce a
variety of carotenoids. We examined a carote-
noid naturally abundant in North Carolina swine
waste lagoons dominated by PNSB. Analytical
methods including high performance liquid ch-
romatography (HPLC), mass spectrometry, and
nuclear magnetic resonance (NMR) confirmed
the identity of the dominant carotenoid as spi-
rilloxanthin, C42H60O2, with 13 conjugated double
bonds. This structure confers antioxidant prop-
erties as good as those of carotenoids currently
marketed as antioxidants. Visual estimates of
the “redness” of swine waste lagoon liquids
were highly correlated with carotenoid content.
Spirilloxanthin concentrations in a lagoon with a
strong PNSB bloom were approximately 0.5
grams·m−3. These results support further inves-
tigations into the potential for extracting com-
mercially valuable natural products from swine
waste lagoons.
Keywords: Swine Waste; Purple Ph ototrophic
Bacteria; Carotenoids; Spirilloxanthin
1. INTRODUCTION
Intensive swine production has created several sig-
nificant environmental challenges, particularly through
widespread use of anaerobic waste storage lagoons and
land application of lagoon liquids. This waste treatment
approach can cause air quality problems with odors [1],
gaseous ammonia emissions [2], greenhouse gas emis-
sions (particularly methane), and airborne transport of
particulates and aerosols [3], surface water quality prob-
lems via lagoon breaches [4], nutrient export from spray
fields [5] and downwind wet and dry aerial deposition
[6], and groundwater contamination, particularly by ex-
cess nitrate [7]. Disputes over these issues in North
Carolina led to an agreement among government, indus-
try, and university researchers to seek alternative waste
treatment systems, leading to development of several
Environmentally Superior Technology (EST) methods [8,
9] and subsequent second generation versions of these
methods [10]. Similar challenges are being addressed by
researchers elsewhere [11-13].
The primary consideration in development of ESTs is
minimization at economically feasible costs of environ-
mental problems associated with conventional waste
treatment methods. Most alternative waste treatment
systems focus on methods of recovering bulk commodi-
ties, e.g., nutrients and bio-solids, or of converting waste
into usable sources of energy, directly through methane
generation and capture or via biological and chemical
transformations into biofuels [14]. Economic benefits to
swine producers were also an important consideration;
North Carolina’s process of EST development has not yet

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815 807
created an economically viable alternative to conven-
tional methods. Moreover, the products of these EST
systems have relatively low market value, owing partly
to low prices for these commodities, geographically con-
centrated production in excess of local demand, and rela-
tively high costs of transport to export markets. Conse-
quently, in 2007 the North Carolina legislature stipulated
use of EST waste treatment systems by new or expanded
swine production operations, but left intact the use of
conventional lagoon-spray field waste treatment systems
by the approximately 2300 existing swine concentrated
animal feeding operations (CAFOs) in North Carolina
[15]. The problem with North Carolina’s approach may
have been, as Kronenberg and Winkler [16] argued, a
focus on process rather than outcome.
We consider here the potential for utilization of natu-
rally dominant microflora in conventional swine waste
lagoons as potential sources of valuable natural products
[17]. Swine waste management experts have recognized
the importance of microbial assemblages in anaerobic
waste treatment, and have provided recommendations for
loading rates that support dominance by the purple non-
sulfur bacteria (PNSB) that metabolize swine waste con-
stituents particularly well [18-20]. These bacteria are
metabolically versatile [21], are capable of photosyn-
thetic and heterotrophic production, use reduced com-
pounds, including many odorants [20,22], as electron
donors, and biosynthesize a variety of potentially useful
natural products [23]. We focus on one natural product, a
carotenoid, from the PNSB that naturally dominate a
significant number of swine lagoons in North Carolina,
conferring a pink-purple color to them at least seasonally.
The pink-purple color of this PNSB assemblage is con-
ferred primarily by carotenoids of the spirilloxanthin and
lycopene synthesis series [24,25]. Lycopene is currently
marketed as a dietary supplement for its potent anti-
oxidant properties [26]. The primary chromophore of
these carotenoids is a long conjugated double bond sys-
tem (11 - 13 carbon-carbon double bonds), which also
confers antioxidant properties of potential commercial
interest [27]. We report here the isolation, identification,
quantification, and characterization of a carotenoid com-
pound, spirilloxanthin, from swine waste lagoons in
southeastern North Carolina.
2. MATERIALS AND METHODS
2.1. Sample Collection
Swine lagoon samples were obtained in bulk quantities
(5 - 10 L) for identification of the dominant carotenoid as
needed from a cooperating swine producer in Pender
County, NC, whose lagoon was dominated by PNSB
year-round. Samples were retrieved in sterile, 1-L screw
cap bottles and stored in refrigerators at 4˚C. Experience
showed that PNSB concentrations were stable for 1 - 2
months under these conditions, and that the bacteria
would float to the top over time, facilitating their re-
moval for analytical work. Lagoon liquid is a complex
matrix of bacteria, other organisms, other particulate
matter, and numerous dissolved constituents. The domi-
nant purple phototrophic bacteria in the samples ana-
lyzed contained substantial amounts of carotenoids, bac-
teriochlorophylls, and other lipid compounds that re-
quired development of extraction and separation tech-
niques suitable for this material. Consequently, identifi-
cation of the principal carotenoid in waste lagoon liquid
employed extraction and liquid chromatographic separa-
tion procedures followed by mass spectrometry and nu-
clear magnetic resonance (NMR) analyses. Additional
sets of swine waste lagoon samples were obtained
through North Carolina State University’s Animal and
Poultry Waste Management Center from swine producers
and from a cooperating swine integrator company on
several occasions as part of their regular quarterly waste
sampling and analysis program using sterile 0.5 L bottles.
These samples were also refrigerated until sub-samples
were withdrawn (after mixing the contents thoroughly)
for use in analyses comparing various properties among
lagoons.
2.2. Sample Preparation
Samples for analysis of total solids concentrations
were mixed thoroughly before decanting known volumes,
freezing, lyophilizing, and weighing dried solids to ±1
mg on a Denver Instruments A-160 electro-balance.
Floating material with high contents of purple phototro-
phic bacteria was decanted from sample bottles, frozen at
−80˚C, and lyophilized using a Virtis Benchtop Freeze
Drier. The resulting purple powder was stored at −4˚C.
The initial extraction protocol utilized a modified caro-
tenoid extraction procedure from De Leenheer and Nellis
[28], in which 3 - 4 washes of material with a mixture of
reagent-grade KOH (60%) and methanol (MeOH) (1:10
ratio by volume) were used to saponify and remove lip-
ids and bacteriochlorophylls. Following centrifugation,
the supernatant was decanted and the remaining solid
material was washed 2× with water, then extracted for 2 -
4 hours in 100% high performance liquid chromatogra-
phy (HPLC) grade acetone. This extract was then filtered
(0.2 µm) and cleaned by solid phase extraction (SPE).
The filtered acetone extract was diluted to ~80% with
deionized water, then drawn through a Fisher Sep-Pak
reversed phase C18 SPE column that had been condi-
tioned with 100% acetone and water. Retained carote-
noids yielded an orange-red color in the column packing.
Following a wash step with 100% methanol, carotenoids
were eluted with a 60:40 mixture of HPLC-grade methyl

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815
808
tert-butyl ether (MTBE) and MeOH. These cleaned ca-
rotenoid extracts were used immediately in HPLC sepa-
rations and analyses.
2.3. Analytical Methods-Carotenoid
The method of Niedzwiedzki et al. [29] was used with
modifications to prepare pigment samples for mass spec-
tral analysis. Cells were mixed with 30 ml of MeOH/
acetone 1:1 and centrifuged in Corex glass tubes at 6000
g for 5 min in a Sorvall SS34 rotor. The supernatant con-
taining a mixture of bacteriochlorophyll and carotenoids
was collected and dried using a rotary evaporator. The
dry residue was dissolved in ~200 ml of MeOH/petro-
leum ether (9:1) and saponified using 5% w/v KOH to
decompose the bacteriochlorophyll. The solution was
then washed with water in a separatory funnel and the
ether layer containing the carotenoids was collected and
dried using a stream of nitrogen gas. The final step was
chromatography over silica using 85% hexanes/15%
acetone as the isocratic mobile phase. The sample eluted
in bands of distinct colors, with the first fraction being
dark orange. This fraction was stored at −80˚C under N2
for subsequent mass spectral analysis. Alternately, the
method of Komori et al. [30] was used with modifica-
tions for mass spectral sample preparation. The bulk of
the bacteriochlorophyll was removed by a single extrac-
tion with ~200 ml MeOH. Dehydrated pellets were ex-
tracted 3X with 200 ml acetone. N-hexane (1 volume)
and water (1 volume) were added to the pooled acetone
extract (1 volume) to transfer the pigments to the n-
hexane layer. The n-hexane fraction was dried in vacuo.
The final step was chromatography over silica, as above.
Samples for NMR analyses were prepared according
to [28] with modifications. Liquid waste was sampled
from the lagoon and placed in volumetric flasks for sev-
eral days. The pinkish material that floated to the top was
lyophilized. In a typical procedure about 1 g of the ly-
ophilized powder was treated with 60% KOH for two
hours in the dark with occasional swirling. The solution
was diluted six-fold with water and then centrifuged at
16,000 - 17,000 rpm in a Beckman JA-20 rotor for 45 -
60 minutes at about 10˚C. The supernatant was discarded
and the jelly-like pellet was extracted with 10 ml of
phenol using a tissue homogenizer. The phenol solution
was extracted using a 50/50 mixture of dichloromethane
and cyclohexane as the organic phase and 10% KOH/5%
NaCl as the aqueous phase. If an emulsion formed it was
broken by filtration using celite. The organic layer was
dried with MgSO4 and the solvent was removed by ro-
tary evaporation. The solid was taken up in approxi-
mately 25 ml of hexane and 10 ml of MeOH and trans-
ferred to a separatory funnel and shaken. The methanol
layer was extracted twice more with portions of hexane.
The combined hexane layers were extracted with a fresh
portion of MeOH, and the second portion of MeOH was
extracted with 2 - 3 portions of hexane. Hexane was re-
moved with rotary evaporation.
The final purification step was reversed phase HPLC
(RP-HPLC) using a Hewlett Packard HP 1100 system
with photo-diode array UV-Vis detection, column tem-
perature set at 25˚C, and a semi-preparatory C-30 col-
umn (NEST group; a Maccel 150 × 10.4 mm 200-5-C30
column, 5 µm particle size and 200 Å pore size). The
mobile phase was 60% MTBE and 40% MeOH, and the
flow rate was 5 ml per minute. The MTBE had been
treated with alumina to remove peroxides then decanted
and filtered as usual. The solid was dissolved in the mo-
bile phase plus 10% CH2Cl2, passed through a syringe
filter, and loaded onto a 2.0 ml sample loop. Compounds
were detected by monitoring at 210, 495, and 700 nm
using diode array detection. The last compound to elute
was the carotenoid of interest, although at least four
other carotenoids were present in smaller amounts. The
volume of the combined fractions was noted, and the
absorbance spectrum (300 - 700 nm) was taken to esti-
mate the amount of carotenoid. The column was peri-
odically cleaned using a gradient of MeOH and chloro-
form, with an additional cleaning step using hexane also
performed in some instances. Smaller samples were
typically purified using a 150 × 4.6 mm column of the
same stationary phase at a flow rate of 1.0 ml/min.
Sample fractions corresponding to the dominant caro-
tenoid peak fraction were analyzed by mass spectrometry
and NMR. High resolution mass spectrometers were
used in analyses of separate carotenoid fraction samples:
an Applied Biosystems Q-trap 2000 LC-MS system at
UNC Wilmington’s Center for Marine Science, and a
JEOL JMS-SX102A HRMS system and an Agilent 6224
LCMS-TOF system with an Agilent series 1200 LC at
the Department of Chemistry at Duke University. NMR
data were acquired on a Varian Inova 800 MHz NMR
equipped with a cryoprobe at the Duke Magnetic Reso-
nance Spectroscopy Center. Standard 1D 1H, 1H/1H
COSY (correlated spectroscopy), 1H/1H TOCSY (total
correlation spectroscopy), 1H/13C HSQC (heteronuclear
single quantum coherence), and 1H/13C HMBC (hetero-
nuclear multiple bond coherence) spectra were obtained
at room temperature in 99.96% CDCl3 (Cambridge Iso-
tope Laboratories, Andover, MA). The program
WINDNMR-pro (Hans J. Reich) was used for spectral
simulations.
2.4. Comparisons among Lagoon Samples
Properties of samples from a variety of swine waste
lagoons analyzed and reported include total carotenoid
and bacteriochlorophyll contents and visible color spec-

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815 809
trum. Total carotenoid content in swine waste samples
was measured as absorbance at 495 nm in acetone ex-
tracts of materials filtered onto Gelman A/E glass fiber
filters. Typical filtered volumes were 2 - 4 ml/sample.
Bacteriochlorophyll content was measured as absorbance
at 780 nm for the same extracted samples.
The visible color of swine waste lagoon samples was
analyzed using a digital (2 MB) photograph of each
waste lagoon sample in a clear polystyrene Petri plate.
Each of the images (124 total) was produced identically
in focus, size and resolution with a custom camera mount,
and included a white/gray/black color target to allow
exact color temperature matching among the individual
photographs. Each image was sampled and ordinated
according to its measured hue value (most red to least
red) expressed in degrees (0˚ - 360˚) in the standard HSV
(Hue, Saturation, and Value or Brightness, HSB) color
space. The red primary area in the color space was ap-
proximately 350˚ to 10˚. The color study representation
may be seen here: http ://gallery.me.com/williamfridrich#
100282. For information on HSV see: http://en.wikipedia.
org/wiki/HSL_color_space.
2.5. Antioxidant Capacity Analyses
The anti-oxidant properties of the principal carotenoid
isolated from swine lagoon waste were further analyzed
using two different published assays for lipophilic anti-
oxidants [31,32]. Purified fractions of swine waste caro-
tenoid were prepared by HPLC using a semi-preparatory
C30 column and isocratic 60:40 MTBE:MeOH elution as
above. These purified fractions were assayed using the
ABTS procedure of Re et al. [31] and the DPPH method
of Brand-Williams et al. [32] with some modifications.
ABTS (2,2’-azinobis(3-ethylbenzothiazoline-6-sulfonic
acid) diammonium salt; Sigma) was dissolved in water to
7 mM concentration (5 tablets of ABTS as supplied dis-
solved in 13 ml DI water). Potassium persulfate solution
(2.4 ml @ 2.45 mM) was mixed with ABTS solution and
allowed to react for 24 h in the dark to form a stable,
highly colored free radical solution. To prepare the final
ABTS solution for the assay, 1 ml of ABTS was diluted
with 19 ml ethanol to obtain an absorbance of approxi-
mately 0.70 at 734 nm. For each reaction 150 μl of stan-
dard carotenoid (lycopene and beta-carotene) or swine
waste carotenoid solution was added to 2.85 ml of the
final ABTS solution. The absorbance at 734 nm was
taken using a spectrophotometer each minute for 6 min.
The absorbance values decreased for the duration of the
reaction due to reduction of the ABTS pre-formed radical
cation by hydrogen donation of the antioxidant com-
pound in question. As ABTS is a decolorization assay,
the degree of color reduction during the reaction corre-
sponded with the antioxidant power of the carotenoid.
Molar concentrations of respective carotenoids were de-
termined by published values for molar absorptivities in
known solvents [33]. Decolorization was expressed as
change in absorbance nmol−1 of carotenoid, with higher
values corresponding to greater antioxidant capacity.
DPPH (2,2-diphenyl-1-picrylhydrazyl) is another stable,
highly colored free radical in solution. A stock solution
was prepared by dissolving 24 mg DPPH in 100 ml
MeOH and stored at −10˚C. The working solution was
acquired by adding 10 ml of stock solution to 45 ml
MeOH to obtain an absorbance reading of approximately
1.1 at 515 nm. For each reaction 150 μl of standard caro-
tenoid or swine waste carotenoid was added to 2.85 ml of
the DPPH solution and allowed to react for 24 h in the
dark. Absorbance readings were then measured at 515
nm. Subsequent to the reaction, the absorbance values
decreased due to the reduction of the DPPH free radical
by the donation of hydrogen ions from the carotenoid.
Molar concentrations of respective carotenoids were de-
termined as above, and antioxidant capacity of each ca-
rotenoid was expressed as change in absorbance nmol−1.
2.6. Bacterial Community Composition
Bacterial composition in a lagoon slurry was deter-
mined with triplicate samples collected from the Pender
County lagoon. A serial dilution from 1:1 to 1:1000 was
made from the swine lagoon slurry and 50 µl of each
dilution was spread onto R2A agar plates in duplicate.
The plates were placed in a 30˚C incubator and allowed
to grow for 3 days. Isolates that resulted in pigment for-
mation were transferred to fresh R2A plates and incu-
bated for 24 h. Genomic DNA from the hog lagoon iso-
lates was extracted using the Gentra Puregene DNA
Purification Kit (Gentra Systems, Inc.; Minneapolis,
MN). 16S rRNA genes were amplified using 27F and
685R primers [34] and Promega GoTaq Master Mix
(Promega Corporation; Madison, WI) according to ma-
nufacturer’s protocol with a 55˚C primer annealing in a
50 µl reaction volume. Polymerase chain reaction (PCR)
amplicons with the 1542 base pair (bp) size were deter-
mined by agarose gel electrophoresis (1% (wt/vol)) and
were purified using the Promega Wizard SV Gel and
PCR Clean-Up System (Promega Corporation; Madison,
WI). The purified products were sequenced using the
ABI Prism® 3100 Genetic Analyzer (Applied Biosystems;
Foster City, CA). The sequences obtained were BLAST-
searched (http://www.ncbi.nih.gov/) to determine the
closest matched bacterial species.
3. RESULTS
3.1 Bacterial Community Composition
Bacterial isolates from the swine lagoon samples were

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815
810
selected based on the differences in colony colors and
shapes. Fifty three isolates were obtained and identified
using 16S rRNA gene sequences (Ta b l e 1 ). Twenty two
isolates produced visible pigments based on the growth
on R2A plates. Among them, 7 isolates generating red
pigments belong to the genera of Rhodococcus, Rhodo-
bacter, Pseudomonas, or Thermomo nas (Tab le 1), which
include species capable of synthesizing spirilloxanthin
[24].
Tab le 1. Bacterial isolates from swine waste lagoons and taxo-
nomic identification based on 16S rRNA gene sequences.
Bacterial Isolate Colony Color Bacterial Identification
HL1/HL7/HL9 Red Thermomonas sp.
HL 10 White Nocardiopsis sp.
HL11 White Bacillus sp.
HL 13 Orange Deinococcus sp.
HL14/HL41 Cream Pseudomonas sp.
HL15/HL28/HL45 White Acinetobacter sp.
HL16 Cream Psychrobacter sp.
HL17/HL39/HL43 Yellow Flavobacterium sp.
HL19 Cream Proteus sp.
HL2/HL18 Yellow Sphingobacterium sp.
HL20 Clear Corynebacterium sp.
HL21 White Imtechium sp.
HL22/HL31/HL48/
HL49/HL50/HL51 Yellow Comamonas sp.
HL27 White Rhizobium sp.
HL29 Yellow Chryseobacterium sp.
HL3/HL4/HL8/HL30 White Alcaligenes sp.
HL32/HL36/HL53 Clear Hydrogenophaga sp.
HL33 Yellow Arcobacter sp.
HL34/HL40/HL42 Clear Azonexus sp.
HL35 Clear Acidovorax sp.
HL37 Red Rhodobacter sp.
HL38 White Dechloromonas sp.
HL44 White Porphyromonadaceae sp.
HL46 Green Acidovorax sp.
HL47 Red Pseudomonas sp.
HL5/HL6 Red Rhodococcus sp.
HL52 Yellow Kocuria sp.
3.2. Carotenoid Identification
HPLC separation of swine waste bacterial pigments
yielded chromatograms that typically exhibited a rapidly
eluted peak identified by its absorbance spectrum as
bacteriochlorophyll, followed by a series of peaks with
absorbance spectra consistent with carotenoids. The last
eluting and usually largest peak had an absorbance spec-
trum in HPLC eluent that closely matched literature val-
ues for spirilloxanthin (Figure 1, Table 2).
Mass spectrometry of an HPLC eluent fraction corre-
sponding to the largest HPLC peak using the Applied
Biosystems Q-trap 2000 LC-MS system yielded peaks
with mass estimates of 596.8 and 597.7 amu and tenta-
tive molecular formula of C42H60O2, both consistent with
published values for spirilloxanthin [36]. DART-MS
spectra obtained from a carotenoid fraction that had been
purified over silica yielded an exact mass of 596.4585
amu, which is 0.5 ppm from the calculated mass
596.4588 amu for C42H60O2. The result using EI on the
JEOL SX-102 was also within the 5.0 ppm criterion for
accurate mass confirmation. These results are consistent
with spirilloxanthin. Two other carotenoids were also
observed, with masses of 490.3807 and 613.4617 amu,
but were not further characterized.
Initial 1H NMR assignments were made on the basis of
coupling constants. Initial 13C NMR assignments were
made on the basis of HSQC data. COSY and HMBC
spectra were used to resolve ambiguities, and the latter
spectrum was also used to assign quaternary carbon at-
oms. H-14 was assigned on the basis of spectral simula-
tions of AA’XX’ systems, with the parameters JXX’ = 15
Hz, JAX = 11.2 Hz, and JAX’ = −3 Hz reproducing the
main features of the relevant portion of the spectrum.
The 3JHH coupling constants are J2,3 = 7.6 Hz, J3,4 = 15.6
Hz, J6,7 = 11.3 Hz, J7,8 = 14.9 Hz, J10,11 = 11.4 Hz, and
Figure 1. Absorption spectrum of purified carotenoid in 60:40
MTBE:MeOH HPLC eluent.

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815 811
Tab le 2 . Absorbance spectra (maxima, nm) of HPLC candidate
peak in Figure 1 vs published values for spirilloxanthin [35].
Solvent system pk 1 pk 2pk I pk II pkIIIIII/II
Published values
Methanol 315 385 464 492 52464%
“HPLC eluent” 469 498 53164%
n-hexane (all trans) 365 386 463 492 526
n-hexane (13-cis) 368 386 460 488 520
n-hexane (15-cis) 367 385 462 490 524
This study
60:40 MTBE:MeOH 316.4 386 466.4 494 52862%
J11,12 = 14.8 Hz. The 1H NMR [37] and 1H and 13C [38]
assignments of all trans -spirilloxanthin have been re-
ported. The chemical shifts (Ta b l e 3 ) and coupling con-
stants of our main carotenoid were in close agreement
with these previous reports. A small number of nuclei
could not be assigned independently of the previous as-
signments. The chemical shifts and coupling constants
we observed support the identification of the main caro-
tenoid as being symmetric and in an all-trans configura-
tion. Based on these results, we have identified this ca-
rotenoid as spirilloxanthin (1,1’-Dimethoxy-3,4,3’,4’-te-
tradehydro-1,2,1’,2’-tetrahydro-Ψ,Ψ-carotene; Figure 2).
We note that spirilloxanthin can occur in various cis-
trans isomers, and that it is possible that some of the
earlier peaks in our HPLC chromatograms were spiril-
loxanthin isomers, as their absorbance spectra in HPLC
eluent were very similar.
3.3. Antioxidant Capacity
Antioxidant capacity analyses of the HPLC fraction
identified as spirilloxanthin yielded estimates compara-
ble to lycopene and greater than or equal to β-carotene in
paired comparisons (Table 4). Analysis of variance of
results from ABTS assays revealed no significant differ-
ences among the three carotenoids (F = 0.44, df = 2.37, p
= 0.65). Analysis of variance and Tukey-Kramer a poste-
riori tests for DPPH assays revealed that spirilloxanthin
had antioxidant capacity higher than β-carotene and ca-
pacity equal to that of lycopene (F = 7.11, df = 2.37, p =
0.0024, α = 0.05). Thus, spirilloxanthin had equivalent if
not greater antioxidant capacity than two carotenoid pre-
parations currently marketed as antioxidants.
3.4. Carotenoid Concentration
The approximate spirilloxanthin concentration in the
swine waste lagoon sampled in Pender County was esti-
mated by a combination of methods. Replicate (4×) 45
Figure 2. Spirilloxanthin chemical structure; note 13 conju-
gated double bonds.
Table 3. NMR chemical shifts of putative spirilloxanthin.
Name C-13 chemical shift, ppm H-1 chemical shift, ppm
HCO- 49 3.23
C-1 74.5
C-2 43.9 2.32
C-3 125 5.72
C-4 137.4 6.16
C-5 135.2
C-6 130.6 6.11
C-7 124.8 6.60
C-8 137.4 6.35
C-9 ?
C-10 132.4 6.23
C-11 124.9 6.65
C-12 137.2 6.38
C-13 ?
C-14 132.6 6.27
C-15 130.2 6.65
C-16, C-1724.1 1.16
C-18 13 1.93
C-19 12.9 1.98
C-20 12.9 1.98
Table 4. Results of antioxidant capacity tests of lycopene, β-
carotene, and spirilloxanthin using ABTS and DPPH assays.
ABTS
Carotenoid Lycopene β-carotene Spirilloxanthin
Δ Absorbance nmol−10.932 1.155 1.024
n 10 10 20
s.e. 0.168 0.168 0.119
DPPH
Carotenoid Lycopene β-carotene Spirilloxanthin
Δ Absorbance nmol−10.051 0.039 0.066
n 10 10 20
s.e. 0.0058 0.0058 0.0041

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815
812
ml aliquots of fresh swine waste frozen, freeze-dried,
saponified with methanolic KOH as in Section 2.3 to
remove lipids and bacteriochlorophyll, then extracted
overnight in 10 ml of 80:20 acetone/MeOH. Absorbance
of centrifuged extracts at 496 nm was measured in glass
1 cm cuvets and the concentration of spirilloxanthin was
calculated using the formula: concentration (M) = ab-
sorbance/(molar absorptivity × path length) using a mo-
lar absorptivity value of 147,000 AU mol−1 L cm−1 at 496
nm for spirilloxanthin [39]. Spirilloxanthin represented
about 35% of total peak area at 495 nm for chroma-
tograms of swine waste samples, so the calculated con-
centration was corrected thereby, yielding an average of
0.98 µM spirilloxanthin, or an average of about 0.58
g·m−3 spirilloxanthin (at 596.458 g·mol−1) in raw waste.
Total solids concentrations from this swine waste lagoon
were in the range of ~5 kg·m−3, so spirilloxanthin content
was on the order of 0.01% of total solids by weight. The
presence of other carotenoids could yield total carotenoid
concentrations 2 - 3 times these levels.
Regression analysis compared color wheel scores (ex-
pressed as degrees on a 0 - 360 scale) to carotenoid con-
centrations measured as absorbance @495 nm of acetone
extracts of filtered lagoon samples. This analysis re-
vealed a highly significant relationship, F = 129.8, df =
1.90, p < 0.0001, = 0.59, indicating that a visual
assessment of the “redness” of lagoon color reliably cor-
responded to actual carotenoid content. Lagoons without
significant PNSB blooms frequently appeared dark brown
or pale green.
2
Adj
R
4. DISCUSSION
Swine producers have known for decades that many
waste lagoons designed and maintained with certain
waste loading rates will develop PNSB blooms that help
in odor suppression. Odor management recommenda-
tions have specifically recognized the importance of
waste loading rates and bacterial processes [18]. Conse-
quently, USDA Natural Resource Conservation Service
(NRCS) guidelines for swine waste lagoon establishment
call for a lagoon liquid volume equivalent to at least ap-
proximately 1 cubic ft per pound of live weight of aver-
age-sized animals to be served by that lagoon [40]. In-
dustry experience has shown that this guideline works
well for grower-finisher and sow operations but less well
for nursery operations, likely owing to differences among
swine diets and subsequent waste contents. Using caro-
tenoid concentrations measured chemically and by visual
methods as proxies for PNSB biomass, our data show
that there was considerable variation in PNSB concentra-
tions in swine lagoons in eastern North Carolina. At the
low end of that range were lagoons that were either re-
ceiving no or low waste inputs (secondary lagoons in
sequenced lagoon systems, pale green color) or that may
have been overloaded (dark brown or black colors) and
releasing considerably more odor [41] than lagoons with
high concentrations of bacteriochlorophyll and carote-
noids.
Visual assessments of lagoon color are reasonably ac-
curate predictors of carotenoid content. The heavily col-
ored lagoons whose contents we analyzed likely con-
tained quantities of spirilloxanthin on the order of grams
m−3, although we can not rule out that other heavily col-
ored lagoons may also contain other carotenoids, such as
lycopene, in addition to various carotenoid precursors of
these compounds for total carotenoid concentrations
several times higher. Individual operators can thus easily
assess the status of their lagoons; broader scale assess-
ments can be obtained by use of aerial surveillance.
Google Earth imagery provides informative snapshots of
the relative proportions of the color spectrum represented
by swine waste lagoons across the broader landscape.
The PNSB include species capable of synthesizing the
spirilloxanthin series of carotenoids as well as the lyco-
pene and okenone series, among other carotenoids [42].
We have not identified lycopene or other carotenoids in
any samples analyzed for this work, but have not yet
analyzed samples from sufficient numbers of swine la-
goons to establish predominance by one bacteria/caro-
tenoid type or another, nor have we sampled across suf-
ficient seasons to detect clear temporal variability in ca-
rotenoid production patterns.
The antioxidant properties of spirilloxanthin are con-
sistent with its molecular structure: a relatively long
backbone of 13 conjugated double bonds and very weakly
polar terminal methoxy groups. Two antioxidant capacity
assays demonstrated that spirilloxanthin is similar if not
superior to lycopene and β-carotene in this regard, but
further antioxidant testing would be necessary for de-
velopment of spirilloxanthin as an antioxidant product
for commercial purposes. Carotenoids currently mar-
keted as antioxidants, including lycopene, β-carotene,
astaxanthin, lutein and zeaxanthin, are all derived from
sources in the natural human food chain, whereas spiril-
loxanthin has to the best of our knowledge never been
obtained from human food sources. Consequently, more
detailed evaluations of its properties and possible benefi-
cial uses as a nutraceutical would be required before
commercialization.
Most proposals for alternate approaches to swine
waste management have derived from conventional uses
of animal manures as fertilizer supplements and soil
amendments, traditional agricultural practices. Current
interest in extraction of energy resources from animal
wastes may, in a sense, also derive from traditional uses
of animal feces as fuel, based on high organic content of
most manures. As the Smithfield Agreement process de-

L. B. Cahoon et al. / Agricultural Sciences 3 (2012) 806-815 813
termined in its final evaluation, however, most such ap-
proaches are not economically feasible, as they cost more
to implement than the conventional lagoon-spray field
waste management system and do not yield products of
sufficient value to offset those costs, particularly when
transport costs are considered. The geographic concen-
tration of swine waste production, e.g., predominantly in
eastern North Carolina, creates a locally saturated market
where the value of such products is low, while transport
costs to markets at any distance would be considerable.
The results of this research raise the possibility that
commercially useful natural products may be obtained
from swine waste lagoons, which may be viewed as large
bioreactors. The full extent of the natural products that
may be available from this source is yet to be determined,
as are conditions and lagoon management practices that
may support higher production of desirable products
while performing waste treatment functions adequately.
The full potential of swine waste as a source of microbial
compounds thus remains to be explored. Clearly, further
research is needed to address these questions.
5. ACKNOWLEDGEMENTS
This research was supported by a Research Competitiveness Fund
award from the University of North Carolina General Administration,
by the North Carolina Pork Council, and by funds provided by UNC
Wilmington. We thank the Duke Magnetic Resonance Spectroscopy
Center, which is supported by the NSF, the NIH, HHMI, the North
Carolina Biotechnology Center, and Duke University. We thank our
Pender County collaborator for access to his swine waste lagoon, and
Lynn Worley-Davis and cooperating swine producers for additional
lagoon samples. We thank Nicholas Wilken and Alexander Amaya for
technical assistance.
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