Excerpt from "Assessing the effect of polyculture on the phenological and nutraceutical profile of five microgreen species" isolating the work done on C. ficifolia.

1. Assessing the effect of polyculture on the phenological and nutraceutical profile of five
microgreen species (Cucurbita ficifolia excerpt).
Celis-Rodriguez, Gracia-Soto, López-Ramírez, Marroquín-Rodríguez, Coutiño-Mijangos, Garcia-Lara &
Espinosa-Leal1
Centro de Biotecnología FEMSA, Escuela de Ingeniería y Ciencias, Tecnológico de Monterrey, Campus
Monterrey, Eugenio Garza Sada 2501. Monterrey, N.L., C.P. 64849, México.
1
Correspondence: claudia_espinosa@tec.mx
ABSTRACT
Microgreens constitute a central component of a significant number of rapidly-growing bodies of
horticultural techniques and developments in pharmaconutrition and urban agriculture. However,
despite the associated disciplinary advances, little to no work has been done that falls outside the
scope of mass production for commercial purposes, or that focuses on organisms originating outside
Western Europe and/or the Mediterranean, regardless of nutritional or horticultural potential. The
novel microhorticultural model proposed by this study presents a symbiotic and
micronutrient-complete co-culture model, based on Mesoamerican Milpa systems and growth
practices, with the aim of providing a highly-accessible, democratizable model for addressing food
insecurity and nutritional deficiencies through sustainable household and community cultivation of
fast-growing, highly-desirable indigenous food crops.
Growth was periodically recorded until harvest, and several micronutrients relevant to common
deficiencies were then measured in the resulting greens. The subjects were then planted together in
milpa arrangements, under common growth conditions determined by data from the aforementioned
growth cycle. Content of previously discussed micronutrients was then measured in the milpa-grown
greens and compared, alongside growth velocity, germination percentage, and size metrics, to the
monoculture data.
Statistically-significant differences between monoculture- and polyculture-grown greens were
observed in all measured parameters, including a marked increase in
secondary-metabolite/nutraceutical compound production for all species, with especially large
increases occurring in O. basilicum.
Keywords: microgreens, polyculture, nutraceuticals, phenological, greenhouse.
1

2. 1. INTRODUCTION
Microgreens present a unique avenue for the rapid and effective production of various food crops.[1][2]
Defined as a group of tender and immature crops, with a central stem, two fully developed cotyledon leaves
and one pair of true leaves, have been shown to contain low levels of antinutrients while being rich in
concentrated micronutrients such as terpenoids, polyphenols, carotenoids, and aminoacids.[2][3]
Resulting
nutraceutical applications and flavor-compound concentrations found in microgreens have led to a rapid
increase in public and scientific interest and a rise in new disciplinary developments in microhorticulture.[4][5]
An overwhelming majority of the relevant research has focused almost exclusively on Brassicaceae, thereby
waylaying potentially-relevant species of different geographic origins that can provide distinct nutraceutical
and flavor compounds.[6]
Likewise, most studies and projects emphasize large-scale for-profit culture,
requiring growth conditions achievable mostly or exclusively in laboratory or high-funding commercial
environments, while microgreens can be produced relatively quickly and easily due to low-cost media, small
space requirement, and short production time. It has been reported that some microgreens require 158–236
times less water to grow and to produce the same amount of nutraceutics than their mature counterparts.[7]
Polyculture, also known as interplanting or intercropping, refers to the simultaneous planting of cultivated
species within the same plot of land, allowing for synchronicity of care and harvest.[8]
Intercropping
agrosystems such as the milpa (a co-silviculture model also known as the Three Sisters) provide an efficient,
sustainable, and resilient culture[9][10]
, relying on the symbiotic or competitive relationships formed by
species grown in association in order to provide a reservoir and active exchange of various nutritional
compounds between organisms, thereby augmenting the growth and nutritional content of the harvest, as
well as conserving water and land resources.[11]
Although soilless microgreens and home-grown models have begun to be tested and implemented
internationally,[12]
hydroponic technology requires investment in additional infrastructure such as pumps,
cycling systems, and nutrient solutions.[13]
In the following study, monoculture and polyculture systems were developed in order to test their effects on
5 crop species: Amaranthus cruentus, Cucurbita ficifolia, Medicago sativa, Ocimum basilicum, and Salvia
hispanica, with phenological, nutraceutical, and yield parameters tracked and evaluated.
The establishment of new culture models based on the following results has the potential to introduce novel
ways of increasing micronutritional output under widely-replicable, low-resource conditions.
2

3. 2. MATERIALS AND METHODS
2.1 Plant material (Germplasm and and planting material)
Native, semi-commercial Amaranthus cruentus, Cucurbita ficifolia, Medicago sativa, and Salvia hispanica
seedstock was acquired from LOMBRIZ MX (Zapopan, Jalisco, México), with Ocimum basilicum stock being
procured from Inverfarms (Querétaro, México). All stock was free of pretreatment or modification. Seeds were
stored in sealed, airtight bags at 4°C until ready to plant. Untreated, pulverized “Terra Huerto” brand
(Azcapotzalco, CDMX, Mexico), coco coir was employed as a culture medium.
2.2 Germination tests
2.2.1 Seed pretreatment
Stock was lixiviated for 4 hours in bidistilled sterilized water. The procedure was performed in a laminar flow
cabinet.
2.2.2 Sowing
Three 0.5 L capacity polypropylene containers (16PP, INIX.mx)—hereafter referred to as experimental
units—were employed for all cultures, following a prior disinfection cycle with a 6% NaClO solution and 70%
ethanol, and tap water rinse. The containers were filled with coco coir to a depth of 4cm, where C. ficifolia seeds
were sown at a 30mm depth in concentric circles.
After sowing, the units were watered with 50 ml of local well water, sealed, and placed in a greenhouse
environment (61±5% relative humidity, T = 23±4°C), occluded from all light until first germination was
observed, and watered with 50mL/48h of well water for 15 days until harvest.
The experimental units were monitored daily for 10 days for germination tracking purposes, charting the
species’ germination curve under the aforementioned conditions and calculating germination speed (%/days)
using the Brown-Mayer standard[14]
:
𝐺𝑒𝑟𝑚𝑖𝑛𝑎𝑡𝑖𝑜𝑛 𝑠𝑝𝑒𝑒𝑑 =
𝐺𝐵
− 𝐺𝐴
𝑡𝑓
− 𝑡0
Where Gb and GA are germination percentages on the last and first day of the linear segment of the species’
observed germination curve, and t is measured in days.
2.3 Growth Culture
2.3.1 Monoculture & Polyculture
4 monoculture units were employed per species, following the previously described sowing procedure, along
with 5 polyculture assay units, where C. ficifolia was sown in a small (D = 1 in) ring in the center of the unit.
Figure 1. Monoculture units, C. ficifolia
3

4. Figure 2. Polyculture units 1-3
2.4 Evaluation of phenological characteristics
Microgreens were harvested after the emergence of the first true leaves (t = 15 days). Each plant was removed
from substrate, brushed, rinsed, and photographed for biophysical measurements, obtained from size-adjusted
whole-sample scans run through ImageJ software.
Root:shoot mass proportions were ascertained by cleaving freshly-harvested greens with a scalpel, along the
taproot-stem interface, and weighing the resulting subsamples separately with an analytical scale (Ohaus). The
split samples were weighed first pre-lyophilization and again pre-pulverization in order to determine fresh
weight, dry weight, and final yield, obtained through the following equation:
𝑌(𝑚𝑔 𝐹𝑊/ 𝑐𝑚
2
) =
%𝐺𝐹
*𝑆𝑇
* 𝐴𝐹𝑊
𝐴
Where %GF is the final germination percentage obtained for each species (section 2.2), ST is the total number of
seeds sown, AFW is the average fresh weight of a single microgreen, and A is total substrate surface area per
unit.
2.5 Nutraceutics
2.5.1 Sample preparation
Puente-Garza, et al’s methodology was employed for sample preparation:[15]
whole-plant samples were
harvested, collected, and stored at –80°C overnight before undergoing lyophilization at –50°C, 0.009 mbar.
Lyophilized samples were then pulverized with a mixer-ball molecular mill (MM 400, Retsch, Verder Scientific,
Germany), through a 180-second, 20Hz cycle.
2.5.2 Extraction
2.5.2.1 Polar Extraction
For Phenolic Compound, Antioxidant Capacity and Tocopherol determination, a 50-mg powder sample of each
species was dissolved in 700 μl of 80:20 v:v methanol:water solution spin for 3 seconds in a vortex at 2500 rpm
(Vortex VWR, Standard Heavy-Duty Vortex Mixer) then, shake all tubes for 5 min in vortex at 2500 rpm,
subsequently were placed in a shaker-incubator (Vortemp 1550, Labnet Int. Inc., Edison, NJ, USA) at 500 rpm,
4

5. 25°C for 15 minutes. Afterwards, it was centrifuged (VWR 1814at, Seed Quality Laboratory) at 5000 rpm for 10
minutes. Supernatant was collected and placed in amber 2 ml Eppendorf tubes.[16]
For the ascorbic acid quantification needed to start with 40-mg powder sample of each species according to
Gillespie et al .2007,It was dissolved in 560 μl of 80:20 v:v methanol:water solution spin for 3 seconds in a
vortex at 2500 rpm (Vortex VWR, Standard Heavy-Duty Vortex Mixer) then, shake all tubes for 5 min in vortex
at 2500 rpm, subsequently were placed in a shaker-incubator (Vortemp 1550, Labnet Int. Inc., Edison, NJ, USA)
at 500 rpm, 25°C for 15 minutes. Afterwards, it was centrifuged (VWR 1814at, Seed Quality Laboratory) at
5000 rpm for 10 minutes. Supernatant was collected and placed in amber 2 ml Eppendorf tubes.
2.5.2.2 Non-polar Extraction
A modified Kurilich-Juvik hexane extraction was employed for the isolation of non-polar compounds:[17]
60 mg
per species of pulverized tissue was placed inside 2 ml amber microvials with 600 μl of a 0.1% w/v TBHQ
solution prepared with absolute ethanol. Prepared samples were placed in a shaker-incubator for 5 min at 500
rpm, 85°C. Subsequently, 50 μl of an 80% KOH solution was added. Incubation was repeated, with time
doubled. Post-incubated vials were immediately stored on ice after the addition of 300 μl of bidistilled sterile
water.
Extraction was performed with n-hexane, with 10 min 1300g centrifugation cycles. The aforementioned was
repeated twice where the resulting supernatant from each cycle was transferred to a different vial. Finally, a
wash with 300 μl of bidistilled water was done at 1200g for 10 min where the superior phase was collected in a
different tube.
2.5.3 Nutraceutical Quantification
2.5.3.1 Ascorbic Acid
Total ascorbic acid (AA) was determined based on the method described by Gillespie & Ainsworth et al:[18]
500µL of extract were diluted in 1 mL of 6% TCA; the resulting solution was then diluted 1:20 with 10 mL of
an 80:20, v:v MetOH:water solution.
The sample was then mixed with 50 µL 75 mM phosphate buffer (pH 7.4). Then, 50 µL of 10 mM DTT were
added and incubated at room temperature for 10 min. To remove the excess of DTT, 25 µL of 0.5% NEM was
added and incubated for 30 s. Afterwards, the samples were mixed with 250 µL 10% TCA, 200 µL 43% ,
𝐻3
𝑃𝑂4
200µL 4% - - and 100 µL . The sample was incubated at 37°C for 1 h—thus reducing the
α α´ 𝑏𝑖𝑝𝑦𝑟𝑖𝑑𝑦𝑙 𝐹𝑒𝐶𝑙3
pool of oxidized AA—and 200 µL were transferred to a clear 96-well microplate. Absorbance was read at λ =
525 nm in a microplate reader (SynergyTM HT Multi-Detection; BioTek Inc.,Winooski, VT). Total AA was
calculated with the following equation:
𝑛𝑚𝑜𝑙 𝐴𝐴/𝑤𝑒𝑙𝑙 =
𝐴525
−0.024
0.00321
Results were expressed in mg of ascorbic acid per 100 g of fresh weight (FW).
5

6. 2.5.3.2 Free Phenolic Compound Determination
Puente-Garza, et al’s modified Folin-Ciocalteu quantification method was employed for free phenolics
determination.[19]
20 μL samples were placed in a microplate with 100 μL of 10% Folin reagent and 80 μL of
7.5% w/v Na2CO3; the mixture was incubated for 1.5 hours at 30°C. Absorbance was measured at 𝜆 = 765 nm
using a microplate reader (SynergyTM HT Multi-Detection; BioTek Inc.,Winooski, VT). Results were expressed
as mg of gallic acid equivalents (GAE) per gram dry weight (DW).
2.5.3.3 Carotene Determination
A spectrophotometric determination of total carotene concentration was performed in a microplate reader at 450
nm. The non-polar extracted samples were diluted 1:5 with n-hexane and 200 µL of it were placed in triplicate,
using n-hexane as a blank. To calculate the final carotene concentration of the samples the following formula
was employed:[20]
𝑇𝑜𝑡𝑎𝑙 𝑐𝑎𝑟𝑜𝑡𝑒𝑛𝑒 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (
µ𝑔𝑐
𝑔 𝐷𝑊
) =
10000 * 𝐴𝑏. * 𝑉
𝐸1 𝑐𝑚
1%
* 𝑙 * 𝑔
Where E1%
1cm stands for the molar extinction coefficient of β-carotene in hexane at 450 nm with a value of 2560
(100 mLcm-1
g-1
), Ab. for the absorbance, V the total extraction volume, l is the pathlength (cm) corresponding
to the wells of the microplate.
2.5.3.4 Tocopherol Determination
Tocopherol content (total) was determined using Fabianek, et al’s micromethod for oxidation-based colorimetric
tocopherol determination.[21]
Phosphoric acid-mediated ferrous chloride oxidation was utilized in order to
generate a carotenoid-interference-free solution for use in low-error spectrophotometric determination of total
tocopherols—thus attempting first confirmation of nutritionally-significant tocopherol presence in C. ficifolia
greens (as opposed to previously-ascertained presence in fruits and seeds), and obtaining tocopherol
concentrations for all species. 4 serial dilutions per species of prepared MetOH extract in 700μL of absolute
MetOH were performed (25, 50, 75, and 100μL), to be measured in a SynergyTM HT Multi-Detection (BioTek
Inc.,Winooski, VT) set to 𝜆 = 290nm, d = 1cm.
Final concentration (expressed in mg/100g) was calculated from absorbance using the following equation
(Fabianek’s equation, modified for conversion into wet weight mass concentration), considering the 300µL
initial volume, extracted from 50mg of lyophilized tissue sample:
𝐶𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 (
𝑚𝑔
100𝑔
) =
2.94𝑒
3.13𝑉𝐴
50𝐻
Where V is the volume of extract diluted in the sample, A is the absorbance measurement at 𝜆 = 290nm, 60°C, d
= 1cm, and H is the water percentage in the pre-lyophilization tissue sample.
6

7. 2.5.3.5 Antioxidant Capacity
Antioxidant capacity was determined using oxygen radical absorbance capacity (ORAC) assay. Extracts were
evaluated using a trolox-fluorescein standard (Sigma-Aldrich). Peroxyl radicals were generated by adding
2,2’-azobis(2-amidinopropane) dihydrochloride (Sigma-Aldrich), monitoring the fluorescence loss signal in a
microplate reader (SynergyTM HT Multi-Detection; BioTek Inc.,Winooski, VT). Excitation and emission
absorbance was set at 𝜆 = 485 and 𝜆 = 538 nm, respectively. Results were expressed as µmol of Trolox
equivalents (TE) per gram of dry weight.
2.6 Statistical analysis
Phenological and nutraceutical analyses were subjected to a one-way analysis of variance (ANOVA), and the
means were compared by a Tukey test (p < 0.05). The statistical analysis was performed using Minitab 21.2
Statistical Software.
3. RESULTS AND DISCUSSION
3.1 Germination tests
Germination was tracked for 10 days for all species (although C. ficifolia did not reach its maximum observed
germination percentage until day 12 in the monoculture units):
Figure 3. Germination rate, monoculture
Germination for C. ficifolia was not observable until unsealed and uncovered (after other species assessed had
germinated, t = 5). However, final germination percentages (~99%, median) far surpassed those previously
reported (~86%)[17]
in both monoculture and polyculture assays, with polyculture units experiencing a marked
increase in germination speed (+9.4%/day), corresponding to a 3-day decrease in time to 99% germination and,
therefore, harvest time.
7

8. 3.3 Evaluation of phenological characteristics
Table 1. Phenological parameters.
Parameter Monoculture Polyculture
Hypocotyl
Length (cm) 4.857±0.188 3.467±0.570
Fresh weight (mg) 2.935±0.223 0.643±0.360
% H2O 88.15% 90.72%
Root system
Length (cm) 15.373±3.468 23.897±1.463
Fresh weight (mg) 1.5±0.294 0.22±0.294
% H2O 89.01% 91.83%
Yield (mg/cm2
) 451 526
C. ficifolia saw some of the highest-percentage changes in water content, mass distribution, and root:shoot
length proportions (Table 1.), with competitive pressures[22]
most likely accounting for the observed differences.
However, final shoot size and weight showed little to no statistically-significant difference, with what little
change occurred affecting yield positively.
3.4 Nutraceutical Quantification
Table 2. Nutraceutical profile.
Parameter Monoculture Polyculture Δ P-value
Tocopherol
(mg/100g)
703±0.634 633±0.036 -9.92% 0.018
Carotenoids
(mg/100g)
749±4.26 558±19.94 -25.52% 0.119
Ascorbic
Acid
(mg/100g)
28,261±0.27 44,262±0.63 +56.62% <0.001
Free Phenols
(mg/100g)
0.668±0.120 0.764±0.490 +14.34% 0.017
ORAC
(μmol/100g)
9.313±0.557 6.992±0.254 -33.20% <0.001
8

9. 3.4.1 Ascorbic Acid
The studied specimens were subjected to competitive stress due to either aerial or root competition, which might
have propitiated the formation of reactive oxygen species (ROS) in the affected tissue, thus triggering the
overexpression of antioxidant compounds such as ascorbic acid—a non-enzymatic antioxidant that counteracts
ROSs in plant tissue.[23]
This might partially explain the increase of AA concentration in every species analyzed,
along with its strong correlation with germination promotion and early stages of plantlet development.[24]
The
later fact plays a key role, especially where microgreens are involved.
3.4.2 Total Phenolic Compounds
Quantification of total phenolics showed an increase in all species. This is indicative of a protective stress
response, with phenolic compound functions mirroring the previously-discussed antioxidant functions of
ascorbic acid.[25]
3.4.3 Carotene Determination
Carotene concentration in analyzed C. ficifolia specimens decreased drastically (-25.52%), with a possible
correlation with decreased overall cotyledon size and true-leaf shoot size, leading to reduced
photosynthetically-active tissue, and, therefore, activity.[26]
3.4.4 Tocopherol Determination
Assessed tocopherol concentrations increased for most of species studied, with double-digit increase being the
norm, in keeping with reported links between oxidative stress—which can be brought about as a result of
highly-intraspecific-competitive environments—and in-tissue tocopherol synthesis as a metabolic amelioration
response.[27]
However, despite the overall trend, significant (−9.92%, P = 0.018) decrease in tocopherol
metabolism was observed in C. ficifolia, a phenomenon not previously observed in the scant literature available
concerning non-fruiting tissues of said species.
3.4.5 Antioxidant Capacity
All the species analyzed displayed a higher antioxidant capacity in monoculture, possibly indicating an increase
in stress-induced ROS synthesis that outpaces the organism’s capacity to produce counteractive compounds.
The reviewed literature indicates that polyphenols are present as majoritary antioxidants in C. ficifolia.
However, said antioxidant activity is only observed up to a certain concentration: at a certain amount of
ROS-induced polyphenol production, this capability is fully lost, and phenolic compounds begin to behave as
prooxidants through induced initiation reactions,[28]
causing the plantlet’s antioxidant capacity to decrease
severely.
9

10. 4. Conclusion
Although further research into the exact metabolic mechanism of C. ficifolia’s reaction to polyculture protocols,
its use as a microgreen, and the workings of its non-fruiting tissue overall is heavily recommended, the present
study has hereby ascertained the species’ suitability for low-cost, high-micronutrient cultivation as a
microgreen, and the quantifiable improvement of both its products and its plot-mates’ through co-culture. It has
also, incidentally, confirmed the presence of nutritionally-significant tocopherol content in non-fruit C. ficifolia
tissue.
10

11. 5. References
1. Teng J, Liao P, Wang MF. (2021) The role of emerging micro-scale vegetables in human diet and
health benefits-an updated review based on microgreens. FOOD Funct. 12(5):1914–32.
2. Ebert AW. (2021) Sprouts and Microgreens—Novel Food Sources for Healthy Diets. Plants, 11(4).
3. Galieni A, Falcinelli B, Stagnari F, Datti A, Benincasa P. (2020) Sprouts and Microgreens: Trends,
Opportunities, and Horizons for Novel Research. AGRONOMY-BASEL. 2020;10(9).
4. Kondratenko EP, Vityaz SN, Miroshina TA, Kuznetsov AS. (2022) Microgreens - biologically
complete product of the XXI century. BIO Web Conf. 42:01002.
5. Sharma S, Dhingra P, Koranne S. (2020) Microgreens: Exciting new food for 21st century. Ecol
Environ Conserv. 26:S248–51.
6. Kyriacou, M. C., Rouphael, Y., Di Gioia, F., Kyratzis, A., Serio, F., Renna, M., ... & Santamaria, P.
(2016). Micro-scale vegetable production and the rise of microgreens. Trends in food science &
technology, 57, 103-115.
7. Sharma, S., Shree, B., Sharma, D., Kumar, S., Kumar, V., Sharma, R., & Saini, R. (2022). Vegetable
microgreens: The gleam of next generation superfoods, their genetic enhancement, health benefits
and processing approaches. Food Research International, 111038.
8. Igbozurike, U. M. (1978). Polyculture and monoculture: contrast and analysis. GeoJournal, 2(5),
443-449.
9. Novotny, I. P., Tittonell, P., Fuentes-Ponce, M. H., López-Ridaura, S., & Rossing, W. A. (2021). The
importance of the traditional milpa in food security and nutritional self-sufficiency in the highlands of
Oaxaca, Mexico. PloS one, 16(2), e0246281.
10. Méndez-Flores, OG, Ochoa-Díaz-López, H, Castro-Quezada, I, Olivo-Vidal, ZE, García-Miranda, R,
Rodríguez-Robles, U, ... & Sánchez-Chino, XM (2021). The milpa as a supplier of bioactive
compounds: a review. Food Reviews International, 1-18.
11. Lotero-Velásquez, E, García-Frapolli, E, Blancas, J, Casas, A, & Martínez-Ballesté, A (2022).
Eco-Symbiotic Complementarity and Trading Networks of Natural Resources in Nahua Communities
in Mountain Regions of Mexico. Human Ecology, 50(2), 307-319.
12. Parida, S. (2020). Innovative Farming of Edible Microgreens at Home and their Nutritional
Composition. TEST Eng. and Manag, 83, 17630-17640.
13. Kularbphettong, K., Ampant, U., & Kongrodj, N. (2019). An automated hydroponics system based on
mobile application. International Journal of Information and Education Technology, 9(8), 548-552.
14. Brown, R. F., and D. G. Mayer. (1988) Representing Cumulative Germination. 1. A Critical Analysis
of Single-Value Germination Indices. Annals of Botany 61, no. 2: 117–25.
15. Puente-Garza CA, Espinosa-Leal CA, García-Lara S. (2018) Steroidal Saponin and Flavonol Content
and Antioxidant Activity during Sporophyte Development of Maguey (Agave salmiana). Plant Foods
Hum Nutr 73:287–94.
16. Zavala-López M, García-Lara S. (2017) An improved microscale method for extraction of phenolic
acids from maize. Plant Methods 13.
11

12. 17. Kurilich AC, Juvik JA. (1999) Quantification of Carotenoid and Tocopherol Antioxidants in Zea
mays. Vol. 47, Journal of Agricultural and Food Chemistry. American Chemical Society (ACS). p.
1948–55
18. Gillespie, K. M., & Ainsworth, E. A. (2007). Measurement of reduced, oxidized and total ascorbate
content in plants. Nature Protocols, 2(4), 871–874.
19. Puente-Garza CA, Espinosa-Leal CA, García-Lara S. (2018) Steroidal Saponin and Flavonol Content
and Antioxidant Activity during Sporophyte Development of Maguey (Agave salmiana). Plant Foods
Hum Nutr 73:287–94.
20. Knockaert G, Lemmens L, Van Buggenhout S, Hendrickx M, Van Loey A. (2012) Changes in
β-carotene bioaccessibility and concentration during processing of carrot puree. Vol. 133, Food
Chemistry. Elsevier BV. p. 60–7
21. Fabianek J, DeFilippi J, Rickards T, Herp A. (1968) Micromethod for tocopherol determination in
blood serum. Clinical Chemistry. 14(5):456-62.
22. Geno, LM, & Geno, BJ. (2001) Polyculture production: principles, benefits and risks of multiple
cropping land management systems for Australia: a report for the rural industries research and
development corporation. Rural Industries Research and Development Corporation.
23. Robinson, D., Davidson, H., Trinder, C., & Brooker, R. (2010). Root–shoot growth responses during
interspecific competition quantified using allometric modelling. Annals of botany, 106(6), 921-926.
24. Akram NA, Shafiq F, Ashraf M. (2018) Ascorbic Acid-A Potential Oxidant Scavenger and Its Role in
Plant Development and Abiotic Stress Tolerance. Front Plant Sci. 8.
25. Tuladhar P, Sasidharan S, Saudagar P. (2021) Role of phenols and polyphenols in plant defense
response to biotic and abiotic stresses. Biocontrol Agents and Secondary Metabolites:419–41
26. Ruiz-Sola MÁ, Rodríguez-Concepción M. (2012) Carotenoid Biosynthesis in Arabidopsis: A
Colorful Pathway. Vol. 10, The Arabidopsis Book. BioOne. p. e0158.
27. Munné-Bosch, S. (2005). The role of α-tocopherol in plant stress tolerance. Journal of plant
physiology, 162(7), 743-748.
28. Das K, Roychoudhury A. (2014) Reactive oxygen species (ROS) and response of antioxidants as
ROS-scavengers during environmental stress in plants. Front Environ Sci. 2.
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