Vijay Bhosekar_ Research Article_ Frontiers in Plant Science

ORIGINAL RESEARCH
published: 19 August 2015
doi: 10.3389/fpls.2015.00652
Edited by:
Raul Antonio Sperotto,
Centro Universitário Univates, Brazil
Reviewed by:
Anil Kumar,
G. B. Pant University of Agriculture
and Technology, India
David D. Baltensperger,
Texas A&M University, USA
*Correspondence:
Manish N. Raizada,
Department of Plant Agriculture,
University of Guelph, 50 Stone Road
East, Guelph, ON N1G 2W1, Canada
raizada@uoguelph.ca
Specialty section:
This article was submitted to
Plant Nutrition,
a section of the journal
Frontiers in Plant Science
Received: 27 May 2015
Accepted: 06 August 2015
Published: 19 August 2015
Citation:
Goron TL, Bhosekar VK, Shearer CR,
Watts S and Raizada MN (2015)
Whole plant acclimation responses by
finger millet to low nitrogen stress.
Front. Plant Sci. 6:652.
doi: 10.3389/fpls.2015.00652
Whole plant acclimation responses
by finger millet to low nitrogen stress
Travis L. Goron, Vijay K. Bhosekar, Charles R. Shearer, Sophia Watts and
Manish N. Raizada*
Department of Plant Agriculture, University of Guelph, Guelph, ON, Canada
The small grain cereal, finger millet (FM, Eleusine coracana L. Gaertn), is valued by
subsistence farmers in India and East Africa as a low-input crop. It is reported by farmers
to require no added nitrogen (N), or only residual N, to produce grain. Exact mechanisms
underlying the acclimation responses of FM to low N are largely unknown, both above
and below ground. In particular, the responses of FM roots and root hairs to N or any
other nutrient have not previously been reported. Given its low N requirement, FM also
provides a rare opportunity to study long-term responses to N starvation in a cereal
species. The objective of this study was to survey the shoot and root morphometric
responses of FM, including root hairs, to low N stress. Plants were grown in pails in a
semi-hydroponic system on clay containing extremely low background N, supplemented
with N or no N. To our surprise, plants grown without deliberately added N grew to
maturity, looked relatively normal and produced healthy seed heads. Plants responded
to the low N treatment by decreasing shoot, root, and seed head biomass. These
declines under low N were associated with decreased shoot tiller number, crown root
number, total crown root length and total lateral root length, but with no consistent
changes in root hair traits. Changes in tiller and crown root number appeared to
coordinate the above and below ground acclimation responses to N. We discuss the
remarkable ability of FM to grow to maturity without deliberately added N. The results
suggest that FM should be further explored to understand this trait. Our observations
are consistent with indigenous knowledge from subsistence farmers in Africa and Asia,
where it is reported that this crop can survive extreme environments.
Keywords: finger millet, nitrogen stress, grain, shoot, root, root hair, crown root, lateral root
Introduction
Finger millet (FM, Eleusine coracana L. Gaertn) is one of the small millet cereals (National Research
Council, 1996; Goron and Raizada, 2015), originally native to the Ethiopian highlands (Dida et al.,
2008). FM is largely consumed by marginalized inhabitants of semi-arid Asia and Africa, and
sold to provide subsistence farmers with additional income (Dorosh et al., 2009; Gruère et al.,
2009). FM is also highly valued by local farmers for its ability to grow in adverse agro-climatic
conditions, where major cereal crops such as maize (Zea mays), wheat (Triticum spp.), and rice
(Oryza sativa) fail, and has been noted to tolerate a wide variety of soils (Upadhyaya et al.,
2006).
Abbreviations: DAT, days after transplant; FM, finger millet; NUE, nitrogen use efficiency.
Frontiers in Plant Science | www.frontiersin.org 1 August 2015 | Volume 6 | Article 652

Goron et al. Finger millet under nitrogen stress
Related to the latter point, FM is also valued for its
exceptionally high nitrogen use efficiency (NUE; Gupta et al.,
2014), defined as grain yield per unit of available nitrogen (N;
Moll et al., 1982). Compared to other grain crops such as maize,
FM responds very well to low amounts of N (Roy et al., 2001).
Many subsistence farmers in South Asia have reported to us that
FM can grow without any added N or with only residual N.
A more formal study carried out by the All India Coordinated
Small Millet Improvement Project (AICSMIP) indicated that
while FM responds well to urea N application at 90 kg/ha,
the cost benefit ratio was highest between 0 and 30 kg N/ha1
.
This is a significantly lower N requirement than corn, for
which optimum application rates can be greater than 200 kg/ha
(Debruin and Butzen, 2014). Other work has confirmed these
observations: a four year study in 2011 showed that FM grain
yield stayed consistent between fertilizer application rates of
20–40 kg N/ha (Pradhan et al., 2011). Limited research also
suggests that FM genotypes vary in their NUE (Thilakarathna and
Raizada, 2015), with some genotypes recognized as having higher
responsiveness to applied N (Puttaswamy and Krishnamurthy,
1976; Bhoite and Nimbalkar, 1996; Dubley and Shrivas, 1999;
Gupta et al., 2012, 2013, 2014). There is paucity of literature
concerning development of new FM genotypes with high yield
potential under low N application regimes (Thilakarathna and
Raizada, 2015). However, in recent years more work has been
accomplished in this area (Goron and Raizada, 2015). Breeding
efforts can be facilitated by selection for traits associated
with NUE.
Despite FM being recognized as a high NUE crop (Gupta
et al., 2014), the underlying mechanisms are not well understood.
Crop plants utilize a wide range of acclimation strategies to
mitigate the limitations of low N availability, both morphological
and biochemical, including altering root traits for better
nutrient salvaging, decreasing chlorophyll production, changing
N allocation within the plant, and altering the timing of flowering
(Chapin et al., 1987; Ciampitti et al., 2013).
Particularly poorly characterized acclimation responses to low
N in plants include changes in root growth and architecture.
The cereal root system consists of thick crown roots which
initiate from the base of the stem (the crown region), from
which lateral roots extend and branch; all of these root types can
further initiate root hairs, which are epidermal projections that
assist with nutrient uptake (Figure 1A). Maize exhibits diverse
root responses to low N, for example by increasing the total
length of the root system, decreasing the crown root number,
and increasing the lateral root to crown root ratio (Eghball
and Maranville, 1993; Wang et al., 2004; Chun et al., 2005; Liu
et al., 2009; Gaudin et al., 2011a). To a lesser extent, low N
is associated with altered root hair length and density in crop
grasses (Gaudin et al., 2011a,b). Under extreme N deficiency
(N starvation), Arabidopsis thaliana ecotypes show a range of
root responses to N starvation (Ikram et al., 2012), though such
extreme experiments are more difficult to conduct with the major
cereals, because of their high N requirement for viability after the
seedling stage.
1
http://smallmillets.res.in/html/reports.html
To the best of our knowledge, there have been no reports
concerning root architecture or root hair traits in FM except for
a recent root hair methodology paper by our group (Goron et al.,
2015). Furthermore, we could not find reports in the literature
concerning other detailed morphological acclimation responses
of FM in response to low N. Given its low N requirement and
small seed size as a source of starter N, FM may also provide a
rare opportunity to understand long-term acclimation responses
of a cereal to extreme N stress.
The objective of this study was to survey shoot and
root morphometric acclimation responses of FM to very low
background N. To ensure minimal levels of N, plants were grown
in pails containing an inert clay substrate called TurfaceR
in a
semi-hydroponic system without added N (Tollenaar and Migus,
1984; Figures 1B,C). This system permitted a more detailed
analysis of fine root traits including root hairs, as shown by our
group with maize (Gaudin et al., 2011a,b) and recently in FM
(Goron et al., 2015), compared to excavation from soil. At the
beginning of the study, it was unclear whether FM plants would
reach maturity in the absence of added N.
Materials and Methods
Plant Materials and Growth Conditions
A field experiment was carried out over two growing seasons
(2012 and 2013). A commercial variety of Eleusine coracana
L. Gaertn (FM) seeds was obtained from India. Seeds were
germinated in open trays at room temperature, supplied with
only double distilled water (ddH2O) in a laboratory for 7 days
before transplantation into pails in a field near Guelph, Canada
(43◦53 N, 80◦18 W, 325 m above sea level). Single FM plants
were grown in 22 L plastic pails (28 cm in diameter) containing
TurfaceR
MVP (Profile Products LLC., Buffalo Grove, IL, USA).
This is an inert, baked-clay, coarse growth medium which can
be used in automated field fertigation systems as previously
described by Tollenaar and Migus (1984). Within the field, the
distance between the centers of pails in each row was 35 cm, and
the spacing between rows was 142 cm. Pails were arranged in
blocks consisting of two N treatments (see below). There were
four blocks per row, with blocks distributed in the field across
five rows, for a total of 20 replicates per treatment per season.
Each block was flanked by buffer plants which received varying
amounts of N. Plants were sampled randomly.
Millet plants were irrigated by an automated mechanism
(Figure 1B), zero to three times per day, adjusted throughout the
growing season as required. A concentrated, modified Hoagland’s
solution lacking N was stored in a 340 L reservoir at the field site,
and was diluted to the appropriate concentration by the irrigation
mechanism at a ratio of 1:100 during application. The pH of the
diluted solution was adjusted to 6.5–6.7 by the addition of HCl.
Two fertigation tubes calibrated to deliver a minimum of 10 ml
of nutrient solution per minute were inserted into each pail (Page
et al., 2011). For the positive nitrogen treatment (+N), 1.1 g of
urea was dissolved into 1 L H20 (37 mM total N, to compensate
for leaching) and was provided to the plants at weekly intervals
(13 times total) over the course of the experiment, with control

FIGURE 1 | Guide to the FM root system and the semi-hydroponic field
fertigation growth system used in this study. Diagram of a FM root network
(A). Crown roots are the thick roots that initiate from the crown region at the
base of the shoot. Lateral roots initiate from the crown roots and can
subsequently branch. Root hairs are present on both crown roots (as shown)
and lateral roots. The fertigation growth system employed in this study
consisted of 22 L plastic pails, 28 cm in diameter, filled with an inert baked clay
medium (TurfaceR
; B). Plants were allowed to grow to maturity as shown (C).
Irrigation hoses delivered the nutrient solution except for nitrogen which was
added manually as described.
plants receiving 1 L H2O with no nitrogen (−N) per dose.
Nitrogen (or the water control) was supplied directly to each
experimental unit by hand at the same time of day throughout
the experiment.
Physiological, Seed Head, Shoot, and Root
System Measurements
Chlorophyll levels of shoot tissue from a minimum of five
randomly selected plants in each treatment were obtained with
a Konica Minolta SPAD-502Plus chlorophyll meter (Konica
Minolta, Inc., Tokyo). Similar leaf positions were sampled
between treatments. In 2012, one measurement was made at 106
DAT; in 2013 two measurements were made at 50 and 80 DAT. In
both years, plant flowering was recorded and tracked over time.
A plant was considered to have flowered on the day at which the
inflorescence was first visible. Tiller number was scored at 113
DAT in 2012 and 119 DAT in 2013.
Seed head tissues were harvested for biomass analysis at 134
DAT in 2012 and 135 DAT in 2013. For root and shoot biomass
analyses, at least 10 plants per treatment were collected at 142
DAT in 2012 and 139 DAT in 2013. In 2013, three samples
of each tissue (10 g each of root, shoot, and seed heads) were
dehydrated in a tissue dryer at 82◦C, ground to a fine powder in
liquid nitrogen, and sent to the University of Guelph Agriculture
and Food Laboratory for total N content quantification with
the Dumas combustion method (Fiedler et al., 1973) using a
LECO FP428 nitrogen and protein determinator (LECO Corp.,
St Joseph, MI, USA).
At the time of harvest in both years, five complete root
systems from each treatment were frozen at −20◦C in 50%
ethanol. Before analysis, roots were thawed in water and floated in
30 cm × 42 cm transparent plastic trays. Roots were scanned with
an Epson Expression 10000XL large area scanner (Seiko Epson
Corporation, Suwa, Nagano, Japan), and the resulting images

were analyzed with WinRhizo software (Version PRO2009,
Regent Instruments Inc., Québec, QC, Canada). The analysis
software was set to measure total root length per diameter class
allowing separate quantification of lateral roots (<0.45 mm) and
crown roots (>0.5 mm). Crown root number was scored by
counting visible roots in the crown region.
Root Hair Microscopy and Measurements
Finger millet root hairs were measured using a protocol recently
developed by our group (Goron et al., 2015). Briefly, root systems
were thawed by floating them in water. Four distinct crown
roots of different lengths were selected from each root system
in order to obtain an accurate representation of the range of
crown root ages within each root network. The four different
crown root length classes were kept constant between plants. Five
segments measuring 1 cm in length were dissected from each of
the selected crown roots at equally spaced distances along the
root, and rinsed in double distilled water. Crown root segments
were stained with 0.4% Trypan Blue solution (MP Biomedicals
LLC, Solon, OH, USA) for 10 min, and then rinsed five times
with ddH2O. Root segments were subsequently immersed in 70%
glycerol (Sigma-Aldrich, St. Louis, MO, USA), then examined
with a Leica MZ8 stereomicroscope (Leica Microsystems GmbH,
Wetzlar, Germany) under 5x magnification. Northern Eclipse
software (version 5.0, Empix Imaging Inc., Mississauga, ON,
Canada) was used to capture four non-overlapping graphic
images of each 1 cm root segment with a Sony DXC-950P Power
HAD 3CCD color video camera (Tokyo, Japan). ImageJ software
(Version 1.47, Wayne Rasband, NIH, USA) was used to manually
trace root hairs to quantify root hair length and density by first
calibrating the program to the image of a stage micrometer (1 mm
in length). For root hair length determination, all four images
per crown root segment were used, with up to 10 root hairs in
each image traced. For statistical analyses, the mean root hair
length from all four images was calculated, representing a pool
of up to 40 root hairs. In total, 14,420 root hairs were traced by
hand for the experiment. For root hair density determination,
for each crown root segment, a randomly selected 300 µm sub-
segment was used to count the number of root hairs. For both
root hair length and density measurements, there were a total of
five replicate plants.
Determination of Potential Plant-Available
Nitrogen in the Clay Growth Medium
A description of the physical-chemical properties of Turface
clay is included (Supplementary Table S1). However, as no
data was available concerning the amount of N present in our
clay gravel growth medium, three replicates of 300 g ground
TurfaceR
MVP were sent to the University of Guelph Agriculture
and Food Laboratory for total N content quantification, with
the Dumas combustion method (Etheridge et al., 1998). The
dry clay gravel was found to contain minimal levels of N
(0.053%; Supplementary Table S2). Additionally, to determine N
bioavailability, 170 g of the growth medium was submerged for
24 h in the same N-free nutrient solution provided to plants in
the field. The resulting slurry was filtered to remove the gravel,
and three replicates of the filtrate were sent to the same lab
listed above for total N quantification by the Kjeldahl method
(Keeney and Bremner, 1966). The filtrate was also found to
contain extremely low levels of N (1.42 mg/L total N, equivalent
to 0.1 mM; Supplementary Table S2).
Statistical Analysis
Analyses were performed using GraphPad PrismR
software
(version 6.04; GraphPad Software, Inc., San Diego, CA, USA).
ROUT was used to identify and remove outliers at Q = 1%. There
was concern for the non-random N treatments of the neighboring
border plants, though they were in separate pails. Nevertheless,
to compensate for any position effects of neighboring plants on
the variance of each treatment, non-parametric Mann–Whitney
tests were used to analyze datasets with non-normal values. Non-
normality was identified with Shapiro–Wilk tests where replicate
numbers were sufficient and with Kolmogorov–Smirnov tests
where the sample size was small. Furthermore, unpaired t-tests
with Welch’s correction for unequal standard deviations were
used to generate P-values.
Correlation coefficients were calculated by the two-tailed
Pearson method with a 95% confidence interval. Hypotheses were
evaluated at a type I error rate of 0.10 or 0.05 as indicated.
Results
Finger Millet Survival under N Starvation
To our surprise, plants that did not receive nitrogen (−N)
deliberately at any point during the experiment (over a ∼140 day
period) reached maturity in both 2012 and 2013. The plants
looked normal, but smaller and less bushy, compared to plants
treated with N (+N), and even produced healthy seed heads
(6.5 g per plant in 2012, and 13.7 g per plant in 2013; Figure 2D;
Table 1).
Root, Shoot, and Seed Head Biomass
In both 2012 and 2013, end-season root and shoot biomass
values were significantly lower when plants were treated with −N
compared with +N (Tables 1 and 2; Figures 2 and 3).
However, biomass values varied widely between years, with
2012 plants being notably smaller than 2013 plants, a reflection
of differences in growing conditions between these years. Due
to these differences, as well as differences in methodology
(i.e., recording dry shoot biomass in 2012 and fresh shoot
biomass in 2013), biomass comparisons were restricted to within
each growing season, and no tests of significance across years
were attempted. In 2012 and 2013, plants receiving the −N
treatment showed 27 and 75% declines in seed head biomass,
respectively, compared to plants that had received the +N
treatment (Table 1).
Whole Plant Architecture
Above ground, in both years, the number of tillers per plant
decreased significantly when plants were treated with −N
compared with +N, with a 30% decrease in 2012 and a
60% decrease in 2013 (Table 1). Below ground, complete FM
root systems at maturity showed a high degree of fibrousness

FIGURE 2 | Finger millet shoot responses to low N at different growth stages. FM in 2012 at 7 weeks of growth (A) and at harvest (B). FM in 2013 at
7 weeks of growth (C) and at harvest (D). All pails were 28 cm in diameter.
TABLE 1 | Measurements of above ground traits of FM in response to low nitrogen stress.
Year Treatment Duration Shoot biomass (g)∗ Seed head biomass (g)# Shoot tiller number
2012 +N 142 DAT 92.8 ± 2.23, n = 10 A 8.9 ± 0.43, n = 10 A 4.0 ± 0.26, n = 10 (113 DAT) A
−N 142 DAT 72.3 ± 0.80, n = 10 B 6.5 ± 0.27, n = 10 B 2.8 ± 0.20, n = 10 (113 DAT) B
2013 +N 139 DAT 869.9 ± 53.83, n = 20 a 54.9 ± 6.14, n = 20 a 50.5 ± 1.87, n = 13 (119 DAT) a
−N 139 DAT 168.0 ± 13.42, n = 20 b 13.7 ± 1.45, n = 20 b 19.9 ± 1.51, n = 13 (119 DAT) b
Letters that are different within a year indicate that the means are significantly different (P < 0.05).
∗Dry biomass was measured in 2012, and fresh biomass in 2013.
#Dry biomass was measured both years.
(Figure 3). In 2012, with the smaller plants, no measurements
of root architecture aside from biomass differed between N
treatments (Table 2). In 2013, with the larger, more N-demanding
plants, the number of crown roots on −N plants was 49% less
than +N plants. Total crown root length, total lateral root length,
and total root system lengths were all 37% smaller in −N plants
(Table 2). There was no difference in terms of average crown root
length or the ratio of total lateral to crown root length.
Correlation between Above Ground Biomass
and Root Characteristics
Pearson correlations were performed between above ground
biomass and root characteristics using data collected in 2013
(Figure 4; data from individual plants were not available in
2012 to permit correlations). In 2013, a significant correlation
was observed between the numbers of crown roots and tillers
(Figure 4A). The shoot fresh biomass also significantly and
positively correlated with the crown root number, total crown
root length and total lateral root length (Figures 4B–D,F), but
not with average crown root length (Figure 4E).
Additionally, Pearson correlations were performed
between seed head dry biomass and the aforementioned
root characteristics for the 2013 season. There were positive
correlations between seed head biomass and total root length,
crown root number, total crown root length, and total lateral root
length (Figures 4H–J,L), but not with average crown root length
(Figure 4K), though some correlations were only significant at
P = 0.10 (as indicated).
Combined, these data indicate that the declines in shoot and
seed head biomass observed under severe low-N stress might be
matched by decreases in shoot tiller number as well as the total
lengths of the two main root classes.
Finger Millet Root Hair Quantification
Root hairs were observed to be widespread along the entire
lengths of FM crown roots (Figures 5C,D; Supplementary Figure
S1). There were no consistent significant differences in root
hair length and density in FM in response to N limitation
(Figures 5C,D; Supplementary Figure S1). However, in some
crown root tips, the newly initiating root hairs (on crown root
segment 5) were either significantly shorter and/or less dense
when N-limited (Figures 5C,D; Supplementary Figure S1). Other
trends were observed: root hairs were generally longer and more
dense in the upper (older) portions of the crown roots, whereas
root hairs became shorter and more sparsely distributed toward
the tips of the crown roots (Figures 5C,D). A similar trend was
observed in 2012, though the differences were less pronounced
(Supplementary Figure S1).
Physiological Measurements
In 2013, all plants were examined daily for evidence of flowering,
defined as the point at which the emerging flower was first
observed. Plants given +N began to flower at 85 DAT, while −N
plants did not begin flowering until 89 DAT. In 2012, the FM
plants began flowering on the same day in both treatments.
In both 2012 and 2013, leaf chlorophyll was measured. In
2012, the smaller FM plants showed no significant differences
in chlorophyll between treatments (Table 3). In 2013, with the
larger, more N-demanding plants, leaf chlorophyll (at 50 DAT) of
−N plants was 56% lower than +N plants. At 80 DAT, the −N

TABLE2|MeasurementsofroottraitsofFMinresponsetolownitrogenstress.
YearDurationRootbiomass(g)∗CrownrootnumberTotalcrownrootlength
(mm)
Averagecrownroot
length(mm)
Totallateralrootlength
(mm)
Totalrootlength(mm)Lateralroot:
crownrootratio
2012+N142DAT229.9±13.48,n=10A80.8±2.65,n=5A6490.2±729.71,n=5A79.7±6.87,n=5A20926.6±1752.15,n=5A27415.8±2363.43,n=5A3.3±0.20,n=5A
−N142DAT56.2±6.07,n=10B69.6±7.00,n=5A6365.4±1102.39,n=5A91.9±13.96,n=5A22486.1±2676.98,n=5A28851.6±3748.96,n=5A3.7±0.25,n=5A
2013+N139DAT249.9±37.26,n=15a216.5±27.25,n=5a11134.1±1124.29,n=4a53.2±1.71,n=5a54639.1±6949.32,n=4a65773.2±7957.09,n=4a4.9±0.30,n=4a
−N139DAT136.2±26.25,n=13b110.8±9.05,n=5b6999.8±653.04,n=5b64.1±6.53,n=5a34576.1±4500.85,n=5b41575.9±5123.39,n=5b4.9±0.26,n=5a
Lettersthataredifferentwithinayearindicatethatthemeansaresignificantlydifferent(P<0.05).Displayedvaluesaremean±SEM.
∗Rootfreshweightwasmeasuredbothyears.
FIGURE 3 | Shoot and root responses of FM to nitrogen limitation, at
harvest. Representative mature, intact plants harvested in 2013, provided
with +N and −N treatments (A). The seed heads were pre-harvested. The
white scale bar at the lower right represents a length of 25 cm. Representative
mature, entire root networks harvested in 2012 grown under +N and −N
treatments, left and right respectively (B). The black scale bar at the lower
right represents a length of 10 cm.
treatment had a leaf chlorophyll content that was 23% lower than
+N plants (Table 3). No significant differences were detected
in total N concentration of root, shoot, or seed head tissue
(Table 3).
Discussion
Consistent with observations of subsistence farmers, our results
show that FM is a remarkable crop – surprisingly able to
flower and produce healthy seed heads without any deliberately
added nitrogen (N). In our study, plants were grown in an
artificial baked clay system with very poor N availability. N-poor
environments are typical of the conditions under which this crop
is grown in Sub-Saharan Africa and South Asia where no applied
N or residual N is used (National Research Council, 1996; Goron
and Raizada, 2015). In South Asia, FM is often transplanted from
nurseries (National Research Council, 1996), a practice mimicked
in this study.

FIGURE 4 | Correlations between FM shoot and seed head biomass and
root architecture traits at harvest across nitrogen treatments. Number of
crown roots vs. tillers (A). Shoot fresh weight vs. total root length (B), shoot
fresh weight vs. total crown root length (C), shoot fresh weight vs. number of
crown roots (D), shoot fresh weight vs. calculated average crown root length
(E), and shoot fresh weight vs. total lateral root length (F). Number of crown
roots vs. total crown root length (G). Seed head dry weight vs. total root length
(H), seed head dry weight vs. total crown root length (I), seed head dry weight
vs. number of crown roots (J), seed head dry weight vs. calculated average
crown root length (K), and seed head dry weight vs. total lateral root length (L).
Pearson r values are displayed. A single asterisk denotes signiﬁcance at
P < 0.10. A double asterisk denotes signiﬁcance at P < 0.05.
The growth and acclimation of FM roots and root hairs in
response to nutrient stress have not previously been reported.
In this study, we have surveyed a range of responses employed
by FM for growth in N-poor environments. Consistent with
other cereal crop plants (Hay and Porter, 2006), FM shoot,
root, and seed head biomass decreased in the −N treatment,
associated with declines in shoot tiller number, crown root
number, total crown root length and total lateral root length, but
with no consistent changes in root hair traits. These results are
summarized in a model (Figure 6).

FIGURE 5 | Root hair morphometric traits from 2013 plants at harvest.
In 2013 and 2012 (data in Supplemental Figure S1), four different classes of
crown roots were sampled based on their lengths/ages (A), and then
examined for root hairs at five different segments spaced evenly along each
crown root using light microscopy with 5x magnification (B). Root hairs were
quantified for length (C) and density (D) at different segments of the crown
roots (x-axis) ranging from the top/nearest the shoot (crown root segment 1)
to near the root tip (crown root segment 5). A blue asterisk (∗) directly above
a mean data point denotes a significant difference in the root hair trait (at
P < 0.05) between N treatments within an individual crown root segment.
An asterisk above the green dashed line denotes a significant difference in
the root hair trait (at P < 0.05) between the top crown root segment
(segment 1) and the bottom crown root segment (segment 2) within the +N
treatment (red asterisk) or the −N treatment (black asterisk). All statistical
analyses were performed with unpaired t-tests (with Welch’s correction where
unequal variances required).

Finger Millet may have Efficient Nitrogen
Uptake Ability
The baked-clay gravel (TurfaceR
) in which plants were grown
intrinsically contains very low N (Supplementary Table S2). The
clay gravel used contained 0.053% total N, which is substantially
low compared to the surface layer of most cultivated soils in
which N content can range between 0.06 and 0.5% (Bremner,
1996; Johnston-Monje et al., 2014). Also, it is important to note
that the media utilized in these experiments was not soil, but
hard, inert gravel, from which the majority of nitrogen would be
unavailable for uptake during the life cycle of the FM plants. Total
nitrogen was measured by soaking the clay gravel in the N-free
nutrient solution used in the experiments, and was found to be
1.42 mg/L, or 0.1 mM total N. In rice and maize, these conditions
would be considered to cause N starvation (Schluter et al., 2012;
Yang et al., 2015). To provide perspective, the United States
Environmental Protection Agency defines clean drinking water
for humans as containing no more than 10 mg/L of N as nitrate –
a level more than seven times that which was available to FM
plants over the course of our study2
. Furthermore, our N values
are likely overestimates, as they do not take into account leaching
of N, given that the coarse clay had a low surface area and hence
poor N binding capacity, with large air pockets to facilitate rapid
downward nutrient flow away from the rhizosphere through
holes in the sides of the pails.
The question then arises as to how FM was able to reach
maturity with no deliberately added N, and with such low levels of
N endogenous to the growth system. Furthermore, surprisingly,
plants showed inconsistent differences in chlorophyll between the
−N vs. +N treatments (Table 3). In 2013 the plants receiving
the −N treatment contained significantly less chlorophyll than
the +N plants. This significant difference was not observed
in 2012, although the trend remained similar. Additionally,
2
http://water.epa.gov/drink/contaminants/basicinformation/nitrate.cfm#four
TABLE 3 | Finger millet tissue chlorophyll and nitrogen content in
response to low nitrogen stress.
Year Measurement Treatment
2012 Chlorophyll content
(SPAD)
+N 11.3 ± 1.27, n = 5 (106 DAT) A
−N 8.2 ± 1.43, n = 5 (106 DAT) A
2013 Chlorophyll content
(SPAD)
+N 51.9 ± 4.06, n = 20 (80 DAT) a
−N 39.8 ± 3.07, n = 18 (80 DAT) b
2013 Shoot nitrogen content
(% dry weight)
+N 1.9 ± 0.06, n = 3 (142 DAT) a
−N 2.1 ± 0.19, n = 3 (142 DAT) a
Root nitrogen content
(% dry weight)
+N 1.3 ± 0.31, n = 3 (142 DAT) a
−N 0.9 ± 0.14, n = 3 (142 DAT) a
Seed head nitrogen
content (% dry weight)
+N 2.5 ± 0.17, n = 3 (142 DAT) a
−N 2.1 ± 0.17, n = 3 (142 DAT) a
Letters that are different within a year indicate that the means are significantly
different (P < 0.05).
FIGURE 6 | Summary of the effects of low N stress on FM architecture.
Low N resulted in a reduction in end-season shoot biomass and seed head
biomass associated with fewer tillers compared to plants receiving higher N
(A). Fewer crown roots were initiated in 2013 in response to the −N treatment
(B), likely responsible for the reduction in total lateral root system length (C).
The calculated average crown root length was unchanged between nitrogen
treatments. No consistent changes in root hair length or density occurred for
root hairs along the crown roots in response to the −N treatment (D).
However, in both N treatments, root hairs were generally longer and denser at
the top of the crown roots than at the bottom of the crown roots (D).
there were no significant differences in tissue N concentration
(Table 3). In other cereal crops, N starvation has been shown
to cause consistent declines in chlorophyll (Széles et al., 2012;
Muñoz-Huerta et al., 2013; Wang et al., 2014) and substantial
declines in tissue N concentration (Havlin et al., 1999; Chen
et al., 2015). Here, the plants were grown on an outdoor field in
open pails (Figures 1B,C), closed at the bottom but perforated
with four drainage holes on the side, very close to the bottom
edge (Supplementary Figure S2). We did sometimes observe
roots growing from these drainage holes, but only in older
plants. It is possible that N was taken up by these late stage
roots to assist with grain fill, but nevertheless, the plants would
have needed to grow for months without these hypothetical
scavenging roots, using starter N from an extremely small seed.
Furthermore, we did not observe weeds in the pails (e.g., that

might host N-fixing microbes) nor an obvious mycorrhizal
network at root harvesting and during root imaging. Some N
may have entered from unintended sources such as from animal
activity (we observed bird feathers), N-fixation by lightning,
algae (growth was observed in pots), or microbial growth in the
fertigation lines. These external N inputs were likely miniscule
and suggest that FM roots themselves may have excellent N
uptake ability.
In the future, it may be worth investigating whether FM hosts
N-fixing microbes, as is observed in its grass relative, sugarcane
(Cavalcante and Dobereiner, 1988; James et al., 1994). We
conducted a preliminary experiment to test whether culturable
microbes from FM root and shoot extracts can grow on agar
without N, but no colonies appeared (data not shown).
These results also provide further impetus to examine FM
as a source of novel agronomic traits for genetic dissection at
the molecular level, especially with regards to N uptake and
utilization efficiency mechanisms. For example, the behavior of
Dof1 and Dof2, transcription factors implicated in controlling
many responses to N, have already been characterized in FM
under conditions of varying N applications (Gupta et al., 2014,
2015; Kumar et al., 2014). However, other molecular genetic
studies concerning N in FM are scarce.
In the –N Treatment, Shoot, and Total Seed
Head Biomass Decreases were Associated
with Declines in Tiller Number
Consistent with other cereal crop plants (Hay and Porter, 2006),
FM shoot and total seed head biomass decreased in the –N
treatment (Table 1), which were associated with declines in tiller
number in both 2012 and 2013. Inadequate N supply has long
been known to lower biomass and yield in plants that display tiller
plasticity including wheat, barley, rice, several bioenergy grasses,
and teosinte, the progenitor to modern maize (Mae, 1997; Gaudin
et al., 2011b; Alzueta et al., 2012; Waramit et al., 2014).
In the –N Treatment in 2013, Root Biomass
Declines were Associated with Diminished
Root Traits
In other cereals including maize, low N availability was shown
to increase the length of the root system, decrease the crown
root number, and increase the lateral root to crown root ratio
(Eghball and Maranville, 1993; Wang et al., 2004; Chun et al.,
2005; Liu et al., 2009; Gaudin et al., 2011a). However, in FM plants
harvested in 2013, we observed a decrease in the total length
of both the lateral root system and crown root system in the
absence of N fertilizer, as well as a decrease in crown root number
(Table 2). Lateral root to crown root ratios remained statistically
equivalent between treatments. In 2012 root architecture did not
differ significantly between treatments.
Differences between our observations and the literature might
be explained by the fact that the earlier studies using other
cereals were conducted under conditions of N limitation, while
our FM plants may have experienced extreme N stress (i.e.,
0.1 mM N from the clay growth medium). The majority of
experiments thus far investigating plant root responses to true
N starvation have been conducted in Arabidopsis. A group
of Arabidopsis ecotypes has been identified which respond to
N starvation with decreases in various root traits including
root fresh biomass (Ikram et al., 2012). However, it should
be noted that the majority of these ecotypes responded by
increasing root biomass, indicating there is a substantial degree
of genetic diversity in plant root acclimation responses. Given
these prior results, it will be helpful to phenotype the roots
of multiple genotypes of FM in response to extreme N
stress.
Research into the responses of crop plants to N starvation,
including in maize and barley, is primarily restricted to molecular
analysis, with a focus on the regulation of N assimilation
genes (Lee et al., 1992; Møller et al., 2011; Nacry et al., 2013).
Additionally, because of the difficulties of imposing true N
starvation on nutrient-intensive crops, the stressful condition is
often applied only for a short time, unlike the current study in
which it was the entire duration of the FM life cycle.
Finger Millet Displays Whole-Plant
Coordination of Architectural Traits in
Response to Low N
Tiller number and crown root number were found to be
correlated across the N treatments (Figure 4), indicating that
FM’s above ground and below ground morphologies may be
tightly linked. To ensure that this relationship is robust, similar
correlations should be generated in the future with additional
N treatments (see below). In modern maize, the transcription
factor Teosinte Branched 1 (TB1) is in part responsible for the
repression of tiller outgrowth, and results in the plant’s single-
stem growth habit (Doebley et al., 1995; Lukens and Doebley,
2001). It has been shown that selection pressure during the
domestication of maize from teosinte led to changes in expression
of the Tb1 gene and coordinated declines in tiller and crown root
number (Hubbard et al., 2002; Gaudin et al., 2011b, 2014). As
FM is genetically related to maize, a similar mechanism may be
directing the tight correlation between the number of tillers and
crown roots and it may be of interest to investigate the presence
of Tb1 orthologs in the FM genome.
Crown Root Initiation may be the Primary
Determinant of Changes in Root Architecture
to Support Altered Shoot Growth
Although the relationship between FM yield and above ground
traits including plant height and shoot biomass has been
previously investigated (Wolie and Dessalegn, 2011), this study is,
to the best of our knowledge, the first time that such correlations
have been performed on below ground plant architecture in FM.
The highest correlations observed between FM shoot and seed
head biomass and root traits were with the crown root number,
not the average crown root length (Figure 4). Furthermore,
of the various root traits that changed in response to the
-N treatment, the crown root number showed the greatest
decline (−49%). Total crown root length was found to correlate
tightly with crown root number (Figure 4G). Since lateral
roots originate from crown roots, then combined these results
indicate that declines in crown root initiation in response to
N limitation primarily determine the other root correlations

observed (Figure 4). It seems logical that when plants have
increased access to N, they respond by initiating additional
crown root systems (with their lateral roots and root hairs)
in order to support increased shoot biomass and seed head
production.
The correlations generated here may aid future breeding
efforts and trait association studies, in which below ground
contributors to yield and biomass could be explored. However,
a relatively small amount of data was used to generate these
correlations, and a degree of prudence must be used in the
interpretation of the results. In particular, the consistency of
responses within our +N and −N treatments resulted in
clustered data (Figure 4). In future experiments, it may be useful
to add additional N treatments to make the correlation analyses
more robust.
As more of the FM genome sequence becomes available it
may be helpful to investigate orthologs of genes implicated in
crown root initiation in other cereals including Tb1 (noted above)
and RTCS identified in maize (Hubbard et al., 2002; Taramino
et al., 2007), as well as Crown rootless 1, 4, 5 and WOX11
identified in rice (Inukai et al., 2005; Zhao et al., 2009; Coudert
et al., 2010; Kitomi et al., 2011). Additional candidate genetic
elements include those responsive to auxin or cytokinin signaling,
pathways recognized as contributing to the control of crown root
growth and development (Inukai et al., 2005; Coudert et al., 2010;
Kitomi et al., 2011).
Low N did not Induce Consistent Changes in
Root Hair Length or Density
Based on hand tracing of 14,420 root hairs in this study, we
observed no consistent differences in root hair length and density
in FM in response to the −N treatment (Figure 5; Supplementary
Figure S1). In the literature, the analysis of root hairs in response
to N limitation/starvation in crop plants is understudied, with
contradictory results. Root hairs in Arabidopsis and several
grass species (Holcus lanatus L., Deschampsia flexuosa L., Poa
annua L., and Lolium perenne L.) showed increased length and
density in response to N limitation (Robinson and Rorison,
1987; Bloch et al., 2011). Root hairs were hypothesized to
contain a mechanism for sensing low N stress (Shin et al.,
2005). In maize and teosinte, however, root hairs showed declines
in length and density as a result of low N conditions using
an aeroponics misting system (Gaudin et al., 2011a,b). One
potentially interesting observation from the present study is
that FM root hairs may have been longer and more dense
in response to low N at the tips of some root age classes
(Figures 5C,D; Supplementary Figure S1A). In the future, precise
examination of root hair initiation may help to confirm these
observations.
Study Limitations
To the best of our knowledge, this study is the first attempt
to grow FM in the northern latitudes of Canada. A growing
season which is cooler and shorter than that of FM’s native
East Africa and India (Goron and Raizada, 2015) created a set
of challenges and limitations. For example, it was necessary to
delay germination and transplantation in the spring until local
temperatures could sustain growth. This resulted in important
differences between the 2012 and 2013 growing seasons.
Although the total time between transplantation and harvest was
similar each year (142 DAT in 2012 vs. 139 DAT in 2013), in 2012,
transplantation was performed 18 days later and growth extended
into November which borders the cold, winter season. In 2012,
the late-season conditions became especially sub-optimal for FM
which requires average daily minimum temperatures to be above
18◦C for optimum growth3
; the average temperature on the day of
harvest was −3.3◦C4
. Plants were stunted with a greatly reduced
tiller number in 2012 compared to the 2013 plants (Table 1), and
as a result the plants were from a different growth stage at the time
of harvest (Figures 2B,D). Indeed, many biomass measurements
(Tables 1 and 2) were significantly lower in 2012 than 2013,
although the trends remained similar in terms of the response
to N treatments. Lower biomass in 2012 may also have been
the result of pathogenesis as disease-like spots were observed
at the seedling stage (they completely disappeared within a few
weeks); these symptoms were absent in 2013. As FM is an
indeterminate plant, the early onset of cold in 2012 (compared
to 2013) combined with the slow growth of seedlings (perhaps
due to the disease) might have prevented the initiation of new
tillers and roots in 2012. The cold likely also affected seed heads
(Table 1), as both grain fill and vegetative growth occur in parallel
in FM. Finally, these differences in growing conditions, combined
with differences when measurements were taken relative to the
growth stage between years, may have also contributed to the
inconsistent chlorophyll readings discussed above.
In 2012, the decrease in root biomass from 230 to 56 g in
the −N treatment was not matched by large differences in the
architectural traits (Table 2). In 2012, we were especially cautious
while washing the root systems after harvest to preserve root hairs
and lateral roots. It is likely that fine TurfaceR
clay stuck to the
roots as a result, especially on the larger +N root systems which
were more dense and difficult to wash, resulting in a non-linear
increase in the biomass. In 2013, the roots were washed more
thoroughly.
Another study constraint was that, although the pails used to
grow plants were very large, they may have limited the growth
of the FM root systems, especially in the +N treatment, thus
reducing the significance of the differences between N treatments.
A future experiment involving growing plants directly in field
soil could be informative, but it would not be possible to impose
extreme N stress conditions and would be especially challenging
to phenotype fine root traits.
A final study limitation was that in 2012, shoot dry weight
was measured while in 2013 shoot fresh weight was measured,
creating a challenge to directly compare biomass results between
years (Table 1). As the leaf tissue in grass species is mostly
water (Garnier, 1992; Garnier and Laurent, 1994), the differences
between the shoot biomass data in 2012 vs. 2013 are not
unreasonable. In any case, all comparisons were made within
years to circumvent this issue. For seed heads and roots, however,
consistent measurements were used across both years.
3
http://www.cabi.org/cpc/datasheet/20674#tab1-nav
4
http://climate.weather.gc.ca/

Study Implications and Future Experiments
This study has provided the first detailed description of FM
responses to extreme N stress, including the first description of
root acclimation responses. The results suggest that FM roots
may have extreme N uptake ability. Mapping of the underlying
genes may aid marker assisted selection (MAS) based efforts to
improve NUE in other cereal crops.
Only a single genotype was examined in this study; however,
there are many landraces available for FM from Africa and South
Asia that show diverse adaptations to local climates (Jarvis et al.,
2008; Wolie and Dessalegn, 2011; Goron and Raizada, 2015). It
will be useful to compare the results from this study to a diversity
panel of FM to identify those with particularly high NUE and
uptake ability. Through such guided breeding approaches, it may
be possible to help subsistence farmers in Africa and South Asia
who have poor access to N fertilizer.
Author Contributions
TLG undertook all lab experiments, the 2013 field experiments,
performed all analyses and wrote the manuscript. VKB helped to
conceive the study, and designed and set up the field experiment
in 2012. CRS helped to set up the field experiments and assisted
with plant growth and measurements. SW assisted with root hair
microscopy and morphometric measurements. MNR conceived
of the study and edited the manuscript.
Acknowledgments
We thank Kirit Patel (Canadian Mennonite University,
Canada) for inspiring our interest in FM. We thank Emma
Harris (University of Guelph) for painstaking assistance in
scanning roots. We thank Amelie Gaudin (University of
California, Davis) for valuable advice in root hair scanning
and microscopy. Hugo Gonzalez (University of Guelph) gave
important direction in the operation of the TurfaceR
fertigation
growth system. We thank Clarence Swanton (University
of Guelph) for use of the fertigation system used in this
experiment. We thank Tadros Atalla, Marina Atalla, and Alex
Whittal for assistance with field harvesting. We thank Dylan
Harding (University of Guelph) for reading and editing the
manuscript. TG was supported in part by scholarships from
the University of Guelph, and a QEII-GSST award from
the government of Ontario. This research was supported by
CIFSRF grants to MNR from the Canadian International
Development Research Centre (IDRC) and the Canadian
Department of Foreign Affairs, Trade and Development
(DFATD).
Supplementary Material
The Supplementary Material for this article can be found online
at: http://journal.frontiersin.org/article/10.3389/fpls.2015.00652
References
Alzueta, I., Abeledo, L. G., Mignone, C. M., and Miralles, D. J. (2012).
Differences between wheat and barley in leaf and tillering coordination under
contrasting nitrogen and sulfur conditions. Eur. J. Agron. 41, 92–102. doi:
10.1016/j.eja.2012.04.002
Bhoite, S. V., and Nimbalkar, V. S. (1996). Response of finger millet cultivars to
nitrogen and phosphorus under rain-fed conditions. J. MaharashtraAgric. Univ.
20, 189–190.
Bloch, D., Monshausen, G., Singer, M., Gilroy, S., and Yalovsky, S.
(2011). Nitrogen source interacts with ROP signalling in root hair tip-
growth. Plant Cell Environ. 34, 76–88. doi: 10.1111/j.1365-3040.2010.
02227.x
Bremner, J. M. (1996). “Nitrogen-total,” in Methods of Soil Analysis. Part 3.
Chemical Methods, ed. J. M. Bartels (Madison, WI: Soil Society of America and
American Society of Agronomy), 1085–1091.
Cavalcante, V. A., and Dobereiner, J. (1988). A new acid-tolerant nitrogen-
fixing bacterium associated with sugarcane. Plant Soil 108, 23–31. doi:
10.1007/BF02370096
Chapin, F. S., Bloom, A. J., Field, C. B., and Waring, R. H. (1987). Plant responses
to multiple environmental factors. Bioscience 37, 49–57. doi: 10.2307/13
10177
Chen, Y., Xiao, C., Wu, D., Xia, T., Chen, Q., Chen, F., et al. (2015). Effects of
nitrogen application rate on grain yield and grain nitrogen concentration in
two maize hybrids with contrasting nitrogen remobilization efficiency. Eur. J.
Agron. 62, 79–89. doi: 10.1016/j.eja.2014.09.008
Chun, L., Mi, G., Li, J., Chen, F., and Zhang, F. (2005). Genetic analysis of maize
root characteristics in response to low nitrogen stress. Plant Soil 276, 369–382.
doi: 10.1007/s11104-005-5876-2
Ciampitti, I. A., Murrell, S. T., Camberato, J. J., Tuinstra, M., Xia, Y.,
Friedemann, P., et al. (2013). Physiological dynamics of maize nitrogen uptake
and partitioning in response to plant density and N stress factors: I. vegetative
phase. Crop Sci. 53, 2105–2119. doi: 10.2135/cropsci2013.01.0040
Coudert, Y., Périn, C., Courtois, B., Khong, N. G., and Gantet, P. (2010). Genetic
control of root development in rice, the model cereal. Trends Plant Sci. 15,
219–226. doi: 10.1016/j.tplants.2010.01.008
DeBruin, J., and Butzen, S. (2014). Nitrogen uptake in corn. Pioneer Crop
Insights 24.
Dida, M. M., Wanyera, N., Harrison, M. L., Bennetzen, J. L., and Devos,
K. M. (2008). Population structure and diversity in finger millet (Eleusine
coracana) germplasm. Trop. Plant Biol. 1, 131–141. doi: 10.1007/s12042-008-
9012-3
Doebley, J., Stec, A., and Gustus, C. (1995). Teosinte branched1 and the origin
of maize: evidence for epistasis and the evolution of dominance. Genetics 141,
333–346.
Dorosh, P. A., Dradri, S., and Haggblade, S. (2009). Regional trade, government
policy and food security: recent evidence from Zambia. Food Policy 34, 350–366.
doi: 10.1016/j.foodpol.2009.02.001
Dubley, O. P., and Shrivas, B. N. (1999). Response of finger millet (Eleusine
coracana) genotypes to nitrogen. Indian J. Agron. 44, 564–566.
Eghball, B., and Maranville, J. W. (1993). Root development and nitrogen influx of
corn genotypes grown under combined drought and nitrogen stresses. Agron. J.
85, 147–152. doi: 10.2134/agronj1993.00021962008500010027x
Etheridge, R. D., Pesti, G. M., and Foster, E. H. (1998). A comparison of
nitrogen values obtained utilizing the Kjeldahl nitrogen and Dumas combustion
methodologies (Leco CNS 2000) on samples typical of an animal nutrition
analytical laboratory. Anim. Feed Sci. Technol. 73, 21–28. doi: 10.1016/S0377-
8401(98)00136-9
Fiedler, R., Proksch, G., and Koepf, A. (1973). The determination of total nitrogen
in plant materials with an automatic nitrogen analyser. Anal. Chim. Acta 63,
435–443. doi: 10.1016/S0003-2670(01)82368-0
Garnier, E. (1992). Growth analysis of congeneric annual and perennial grass
species. J. Ecol. 80, 665–675. doi: 10.2307/2260858
Garnier, E., and Laurent, G. (1994). Leaf anatomy, specific mass and water content
in congeneric annual and perennial grass species. New Phytol. 128, 725–736.
doi: 10.1111/j.1469-8137.1994.tb04036.x

Gaudin, A. C. M., McClymont, S. A., Holmes, B. M., Lyons, E., and Raizada,
M. N. (2011a). Novel temporal, fine-scale and growth variation phenotypes
in roots of adult-stage maize (Zea mays L.) in response to low nitrogen
stress. Plant Cell Environ. 34, 2122–2137. doi: 10.1111/j.1365-3040.2011.
02409.x
Gaudin, A. C. M., McClymont, S. A., and Raizada, M. N. (2011b). The nitrogen
adaptation strategy of the wild teosinte ancestor of modern maize, Zea
mays subsp. parviglumis. Crop Sci. 51, 2780–2795. doi: 10.2135/cropsci2010.
12.0686
Gaudin, A. C. M., McClymont, S. A., Soliman, S. S. M., and Raizada, M. N.
(2014). The effect of altered dosage of a mutant allele of Teosinte branched
1 (tb1-ref) on the root system of modern maize. BMC Genet. 15:23. doi:
10.1186/1471-2156-15-23
Goron, T. L., and Raizada, M. N. (2015). Genetic diversity and genomic resources
available for the small millet crops to accelerate a New Green Revolution. Front.
Plant Sci. 6:157. doi: 10.3389/fpls.2015.00157
Goron, T. L., Watts, S., Shearer, C., and Raizada, M. N. (2015). Growth in Turface
clay permits root hair phenotyping along the entire crown root in cereal crops
and demonstrates that root hair growth can extend well beyond the root hair
zone. BMC Res. Notes 8:143. doi: 10.1186/s13104-015-1108-x
Gruère, G., Nagarajan, L., and King, E. D. I. O. (2009). The role of collective
action in the marketing of underutilized plant species: lessons from a
case study on minor millets in South India. Food Policy 34, 39–45. doi:
10.1016/j.foodpol.2008.10.006
Gupta, A. K., Guar, V. S., Gupta, S., and Kumar, A. (2013). Nitrate signals determine
the sensing of nitrogen through differential expression of genes involved in
nitrogen uptake and assimilation in finger millet. Funct. Integr. Genomics 13,
179–190. doi: 10.1007/s10142-013-0311-x
Gupta, N., Gupta, A. K., Gaur, V. S., and Kumar, A. (2012). Relationship of nitrogen
use efficiency with the activities of enzymes involved in nitrogen uptake and
assimilation of finger millet genotypes grown under different nitrogen inputs.
Sci. World J. 2012, 1–10. doi: 10.1100/2012/625731
Gupta, S., Gupta, S. M., Gupta, A. K., Gaur, V. S., and Kumar, A. (2014). Fluctuation
of Dof1/Dof2 expression ratio under the influence of varying nitrogen and light
conditions: involvement in differential regulation of nitrogen metabolism in
two genotypes of finger millet (Eleusine coracana L.). Gene 546, 327–335. doi:
10.1016/j.gene.2014.05.057
Gupta, S., Malviya, N., Kushwaha, H., Nasim, J., Bisht, N. C., Singh, V. K., et al.
(2015). Insights into structural and functional diversity of Dof (DNA binding
with one finger) transcription factor. Planta 241, 549–562. doi: 10.1007/s00425-
014-2239-3
Havlin, J., Beaton, J., Tisdale, S., and Nelson, W. (1999). “Nitrogen,” in Soil Fertility
and Fertilizers, 6th Edn, eds M. Carnis and L. Harvey (Upper Saddle River, NJ:
Prentice-Hall, Inc).
Hay, R., and Porter, J. (2006). “Limiting factors and the achievement of high
yield,” in The Physiology of Crop Yield (Oxford: Blackwell Publishing Inc.),
180–204.
Hubbard, L., McSteen, P., Doebley, J., and Hake, S. (2002). Expression patterns and
mutant phenotype of teosinte branched1 correlate with growth suppression in
maize and teosinte. Genetics 162, 1927–1935.
Ikram, S., Bedu, M., Daniel-Vedele, F., Chaillou, S., and Chardon, F. (2012).
Natural variation of Arabidopsis response to nitrogen availability. J. Exp. Bot.
63, 91–105. doi: 10.1093/jxb/err244
Inukai, Y., Sakamoto, T., Ueguchi-Tanaka, M., Shibata, Y., Gomi, K., Umemura, I.,
et al. (2005). Crown rootless1, which is essential for crown root formation in
rice, is a target of an AUXIN RESPONSE FACTOR in auxin signaling. Plant
Cell 17, 1387–1396. doi: 10.1105/tpc.105.030981
James, E., Reis, V., Olivares, F., Baldani, J., and Döbereiner, J. (1994). Infection of
sugar cane by the nitrogen-fixing bacterium Acetobacter diazotrophicus. J. Exp.
Bot. 45, 757–766. doi: 10.1093/jxb/45.6.757
Jarvis, D. I., Brown, A. H. D., Cuong, P. H., Collado-panduro, L., Latournerie-
moreno, L., Gyawali, S., et al. (2008). A global perspective of the
richness and evenness of tradtitional crop-variety diversity maintained by
farming communities. Proc. Natl. Acad. Sci. U.S.A. 105, 5326–5331. doi:
10.1073/pnas.0800607105
Johnston-Monje, D., Mousa, W., Lazarovits, G., and Raizada, M. N. (2014). Impact
of swapping soils on the endophytic bacterial communities of pre-domesticated,
ancient and modern maize. BMC Plant Biol. 14:233. doi: 10.1186/s12870-014-
0233-3
Keeney, D. R., and Bremner, J. M. (1966). Comparison and evaluation of
labroratory methods of obtaining an index of soil nitrogen availability. Agron. J.
58, 498–503. doi: 10.2134/agronj1966.00021962005800050013x
Kitomi, Y., Ito, H., Hobo, T., Aya, K., Kitano, H., and Inukai, Y. (2011). The
auxin responsive AP2 / ERF transcription factor CROWN ROOTLESS5 is
involved in crown root initiation in rice through the induction of OsRR1,
a type-A response regulator of cytokinin signaling. Plant J. 3, 472–484. doi:
10.1111/j.1365-313X.2011.04610.x
Kumar, A., Kanwal, P., Gupta, A. K., Singh, B. R., and Gaur, V. S. (2014). A full-
length Dof1 transcription factor of finger millet and its response to a circadian
cycle. Plant Mol. Biol. Report. 32, 419–427. doi: 10.1007/s11105-013-0653-5
Lee, R. B., Purves, J. V., Ratcliffe, R. G., and Saker, L. R. (1992). Nitrogen
assimilation and the control of ammonium and nitrate absorption by maize
roots. J. Exp. Bot. 43, 1385–1396. doi: 10.1093/jxb/43.11.1385
Liu, J., Chen, F., Olokhnuud, C., Glass, A. D. M., Tong, Y., Zhang, F., et al. (2009).
Root size and nitrogen-uptake activity in two maize (Zea mays) inbred lines
differing in nitrogen-use efficiency. J. Plant Nutr. Soil Sci. 172, 230–236. doi:
10.1002/jpln.200800028
Lukens, L., and Doebley, J. (2001). Molecular evolution of the teosinte branched
gene among maize and related grasses. Mol. Biol. Evol. 18, 627–638. doi:
10.1093/oxfordjournals.molbev.a003843
Mae, T. (1997). Physiological nitrogen efficiency in rice: nitrogen utilization,
photosynthesis, and yield potential. Plant Soil 196, 201–210. doi:
10.1023/A:1004293706242
Møller, A. L. B., Pedas, P., Andersen, B., Svensson, B., Schjoerring, J. K., and
Finnie, C. (2011). Responses of barley root and shoot proteomes to long-term
nitrogen deficiency, short-term nitrogen starvation and ammonium. Plant Cell
Environ. 34, 2024–2037. doi: 10.1111/j.1365-3040.2011.02396.x
Moll, R. H., Kamprath, E. J., and Jackson, W. A. (1982). Analysis and interpretation
of factors which contribute to efficiency of nitrogen-utilization. Agron. J. 74,
562–564. doi: 10.2134/agronj1982.00021962007400030037x
Muñoz-Huerta, R. F., Guevara-Gonzalez, R. G., Contreras-Medina, L. M., Torres-
Pacheco, I., Prado-Olivarez, J., and Ocampo-Velazquez, R. V. (2013). A review
of methods for sensing the nitrogen status in plants: advantages, disadvantages
and recent advances. Sensors 13, 10823–10843. doi: 10.3390/s1308
10823
Nacry, P., Bouguyon, E., and Gojon, A. (2013). Nitrogen acquisition by roots:
physiological and developmental mechanisms ensuring plant adaptation to a
fluctuating resource. Plant Soil 370, 1–29. doi: 10.1007/s11104-013-1645-9
National Research Council. (1996). Lost Crops of Africa. Washington, DC: National
Academy Press.
Page, E. R., Liu, W., Cerrudo, D., Lee, E. A., and Swanton, C. J. (2011). Shade
avoidance influences stress tolerance in maize. Weed Sci. 59, 326–334. doi:
10.1614/WS-D-10-00159.1
Pradhan, A., Thakur, A., Patel, S., and Mishra, N. (2011). Effect of different nitrogen
levels on kodo and finger millet under rainfed conditions. Res. J. Agric. Sci. 2,
136–138.
Puttaswamy, S., and Krishnamurthy, K. (1976). Relative dry-matter efficiency in
finger millet genotypes in relation to levels of spacing and nitrogen. Mysore J.
Agr. Sci. 10, 345–352.
Robinson, D., and Rorison, I. H. (1987). Root hairs and plant growth at
low nitrogen availabilities. New Phytol. 107, 681–693. doi: 10.1111/j.1469-
8137.1987.tb00906.x
Roy, D., Chakraborty, T., Sounda, G., and Maitra, S. (2001). Effect of fertility
levels and plant population on yield and uptake of nitrogen, phosphorus and
potassium in finger millet (Eleusine coracana) in lateritic soil of West Bengal.
Indian J. Agron. 464, 707–711.
Schluter, U., Mascher, M., Colmsee, C., Scholz, U., Brautigam, A., Fahnenstich, H.,
et al. (2012). Maize source leaf adaptation to nitrogen deficiency affects not only
nitrogen and carbon metabolism but also control of phosphate homeostasis.
Plant Physiol. 160, 1384–1406. doi: 10.1104/pp.112.204420
Shin, R., Berg, R. H., and Schachtman, D. P. (2005). Reactive oxygen species and
root hairs in Arabidopsis root response to nitrogen, phosphorus and potassium
deficiency. Plant Cell Physiol. 46, 1350–1357. doi: 10.1093/pcp/pci145
Széles, A. V., Megyes, A., and Nagy, J. (2012). Irrigation and nitrogen effects on the
leaf chlorophyll content and grain yield of maize in different crop years. Agric.
Water Manag. 107, 133–144. doi: 10.1016/j.agwat.2012.02.001
Taramino, G., Sauer, M., Stauffer, J. L. S. Jr., Multani, D., Niu, X., and Sakai, H.
(2007). The maize (Zea mays L.) RTCS gene encodes a LOB domain protein

that is a key regulator of embryonic seminal and post-embryonic shoot-borne
root initiation. Plant J. 1, 649–659. doi: 10.1111/j.1365-313X.2007.03075.x
Thilakarathna, M. S., and Raizada, M. N. (2015). A review of nutrient management
studies involving finger millet in the semi-arid tropics of Asia and Africa.
Agronomy 5, 262–290. doi: 10.3390/agronomy5030262
Tollenaar, M., and Migus, W. (1984). Dry matter accumulation of maize grown
hydroponically under controlled-environment and field conditions. Can. J.
Plant Sci. 64, 475–485. doi: 10.4141/cjps84-070
Upadhyaya, H. D., Gowda, C. L. L., Pundir, R. P. S., Reddy, V. G., and
Singh, S. (2006). Development of core subset of finger millet germplasm using
geographical origin and data on 14 quantitative traits. Genet. Resour. Crop Evol.
53, 679–685. doi: 10.1007/s10722-004-3228-3
Wang, G., Bronson, K. F., Thorp, K. R., Mon, J., and Badaruddin, M.
(2014). Multiple leaf measurements improve effectiveness of chlorophyll
meter for durum wheat nitrogen management. Crop Sci. 54, 817–826. doi:
10.2135/cropsci2013.03.0160
Wang, Y., Mi, G., Chen, F., and Zhang, J. (2004). Response of root morphology to
nitrate supply and its contribution to nitrogen accumulation in maize. J. Plant
Nutr. 27, 2189–2202. doi: 10.1081/LPLA-200034683
Waramit, N., Moore, K. J., and Heaton, E. (2014). Nitrogen and harvest date affect
developmental morphology and biomass yield of warm-season grasses. GCB
Bioenergy 6, 534–543. doi: 10.1111/gcbb.12086
Wolie, A., and Dessalegn, T. (2011). Correlation and path coefficient analyses
of some yield related traits in finger millet ( Eleusine coracana (L.) Gaertn.)
germplasms in northwest Ethiopia. Afr. J. Agric. Res. 6, 5099–5105. doi:
10.5897/AJAR11.616
Yang, W., Yoon, J., Choi, H., Fan, Y., Chen, R., and An, G. (2015). Transcriptome
analysis of nitrogen-starvation- responsive genes in rice. BMC Plant Biol. 15:31.
doi: 10.1186/s12870-015-0425-5
Zhao, Y., Hu, Y., Dai, M., Huang, L., and Zhou, D.-X. (2009). The WUSCHEL-
related homeobox gene WOX11 is required to activate shoot-borne crown root
development in rice. Plant Cell 21, 736–748. doi: 10.1105/tpc.108.061655
Conflict of Interest Statement: The authors declare that the research was
conducted in the absence of any commercial or financial relationships that could
be construed as a potential conflict of interest.
Copyright © 2015 Goron, Bhosekar, Shearer, Watts and Raizada. This is an open-
access article distributed under the terms of the Creative Commons Attribution
License (CC BY). The use, distribution or reproduction in other forums is permitted,
provided the original author(s) or licensor are credited and that the original
publication in this journal is cited, in accordance with accepted academic practice.
No use, distribution or reproduction is permitted which does not comply with these
terms.

Vijay Bhosekar_ Research Article_ Frontiers in Plant Science

More Related Content

What's hot (19)

Similar to Vijay Bhosekar_ Research Article_ Frontiers in Plant Science (20)

Vijay Bhosekar_ Research Article_ Frontiers in Plant Science