BIOLOGY
Plant Characteristics
St. Augustinegrass
has round-tipped leaf blades 5 to 14 mm wide and they are
arranged in a strictly distichous manner. It is a perennial,
and it spreads by branching stolons, forming a coarse and
spongy canopy. Because of the absence of rhizomes or other
protected stems, it recuperates poorly from defoliation and
has poor wear tolerance. Cultivars with shorter internodes
have higher wear tolerance (Busey, 1991, unpublished data).
Crowns or basal shoot aggregates are absent. The stolon internodes
are exposed, and slightly flattened dorsiventrally.
The leaf blades
are generally glabrous, but genotypes showing possible introgression
with S. dimidiatum (L). Brongn., pembagrass,
are sparsely pubescent (Busey, 1990b). The midrib is conspicuous.
The bases of the leaf blades are attenuated and subtended
by constricted collars, which are conspicuously lighter than
the blade or sheath, making the leaves pseudopetiolate. The
ligule is a minutely ciliate membrane. The leaf sheaths are
compressed laterally, nearly forming a keel. As a C4 species,
St. Augustinegrass has typical Krantz leaf anatomy (Krans
et al., 1979), containing an inner parenchyma bundle sheath
layer with centripetal chloroplasts (Fig. 1). This anatomical
characteristic of C4 grasses facilitates the compartmentalization
of photosynthetic processes in two different cellular regions,
repressing photorespiration (Dengler et al., 1994).
The inflorescences
of St. Augustinegrass are modified spike-like panicles, with
the branches of the inflorescence contracted and often reduced
to single spikelets. Branches are partially embedded in hollows
on one face or the sides of a corky rachis. The rachis, which
is terminated in a naked point, normally disarticulates at
the branch nodes into squarish segments containing the spikelet(s).
The inflorescence segments float in saltwater for 7-10 days,
which may not be sufficient for transoceanic dispersal (Sauer,
1972). The awnless spikelets are 3-6 mm long and have dissimilar
glumes. The lower or first glume is scale-like, only about
1 mm long, and nerveless. The upper or second glume is similar
to, and about the same length as, the nerved, chartaceous
lemmas. Spikelets contain a lower floret, which is most often
staminate or is neuter, but is perfect and sets seed in some
genotypes (Center and Busey, 1981, unpublished data). The
upper perfect floret contains three anthers, which may vary
among genotypes from orange-buff with flecks of purple to
sulfur yellow. The two stigmata may be purple, or translucent
appearing white, or bicolor (purple shafts and translucent
branches). Internodes may vary from purplish to green, in
association with the color of anthers and stigmata. For example,
plants with purplish internodes generally have purple stigmata
and orange-buff anthers, while plants with green internodes
generally have whitish translucent stigmata and sulfur-yellow
anthers.
Environmental
Adaptation and Management
St. Augustinegrass
provides a tight leaf canopy, due to relatively prostrate
leaf angle; therefore, it is highly resistant to weed infestation.
Some cultivars, such as 'Floratam,' grow well in sandy coastal
areas where zoysiagrasses, Zoysia spp., and bermudagrass,
Cynodon spp., grow poorly due to parasitism by the sting nematode,
Belonolaimus longicaudatus Rau (Busey et al., 1982b).
St. Augustinegrass grows adequately across a wider range of
soil conditions than other warm-season turfgrasses. It generally
grows without problems in sand, loam, and humic soils, and
across a pH range from 4.5 to 8.5. Under conditions of high
pH and waterlogged soil, including production in plastic trays,
interveinal chlorosis symptomatic of iron deficiency is sometimes
observed, particularly in the Breviflorus Race (see Taxonomy
and Geography).
Documented long-term
nutritional management studies have not been done. Therefore,
with increasing scrutiny of lawn maintenance practices as
possible nonpoint sources of groundwater pollutants such as
nitrate, the appropriate rates, timings, and nutrition sources
for St. Augustinegrass turf fertilization are unresolved.
St. Augustinegrass is often grown in warm coastal areas with
shallow aquifers, and the appropriate nitrogen fertilization
is therefore especially important in protecting groundwater.
Use of high rates
of inorganic nitrogen has been associated with southern chinch
bug, Blissus insularis Barber, outbreak in St. Augustinegrass
(Busey and Snyder, 1993). Higher fertilization rates are associated
with higher frequency of wilt in St. Augustinegrass turf,
compared with lower fertilization rates (Busey, 1996). Higher
fertilization rates are also associated with higher levels
of thatch, a problem for St. Augustinegrass considering that
it is entirely stoloniferous, and any accumulation of runners
is above ground.
Several cultivars
of St. Augustinegrass tolerate partial shade (Busey and Davis,
1991), a valuable trait for use in lawns, particularly in
smaller residential landscapes and where trees are dominant.
The shade tolerance of St. Augustinegrass is useful in mixed
cropping systems of the tropics, where an herbaceous understory
is grazed by animals, in the diminished illumination beneath
tree crops. St. Augustinegrass has the least reduction in
yield, and the largest yield, among eight grasses evaluated
under the shade of coconuts, Cocos nucifera L., in
the Solomon Islands (Smith and Whiteman, 1983); the coconuts
transmitted 20% relative irradiance (full sunlight=100% irradiance).
Among warm-season turfgrasses, St. Augustinegrass performs
better under reduced illumination than bahiagrass, Paspalum
notatum Flügge; bermudagrasses, Cynodon spp.; centipedegrass,
Eremochloa ophiuroides (Munro) Hack.; And zoysiagrasses,
Zoysia spp. (Beard, 1973).
St. Augustinegrass
generally has 10 to 30% greater evapotranspiration than bermudagrass
in mini-lysimeters under semiarid conditions (Casnoff et al.,
1989; Kim and Beard, 1988; Kneebone and Pepper, 1982). However,
electromagnetic measurement of soil moisture in unrestricted
plot areas under humid conditions showed that the evapotranspiration
of St. Augustinegrass is not significantly different from
bermudagrass (Carrow, 1995). Drought resistance in St. Augustinegrass
is due to drought survival through wilt avoidance due to deeper
or more effective root systems and not by reduced evapotranspiration
(see Physiology and Environmental Stresses). St. Augustinegrass
is a model species for studying water relationships including
evapotranspiration (Stewart and Mills, 1967) and leaf diffusive
resistance (Johns et al., 1983).
DISTRIBUTION,
CYTOTAXONOMY, AND GENETICS
Origin and
Related Species
The genus Stenotaphrum
Trin. is a primarily tropical member of the tribe Paniceae
of the Panicoideae. Whereas S. secundatum, St.
Augustinegrass, occurs on all continents except Antarctica,
the six other species of Stenotaphrum are known naturally
only from East Africa, the islands and coastlines of the Indian
Ocean, and from southern China to the South Pacific (Busey,
1995b; Sauer, 1972). Most occupy restricted natural habitats,
and three species are island endemics. Spikelet and inflorescence
characteristics of Stenotaphrum are most similar to
the monotypic genera Thuarea Pers. and Uranthoecium
Stapf of Australia; Thuarea also occurs in coastal regions
of tropical Asia (Webster, 1988). According to Webster (1988),
the genus Stenotaphrum is probably not closely related
to Paspalidium Stapf, as suggested by Sauer (1972).
Morphologically,
pembagrass, S. dimidiatum, is the species most
similar to St. Augustinegrass; the two species are separated
primarily by number of spikelets per raceme (Sauer, 1972).
Some polyploid St. Augustinegrass introductions show possible
introgression with pembagrass. Inflorescence racemes of the
presumptive introgressants, such as FX-10 and its relatives
(Busey, 1993), produce three and occasionally four spikelets,
which would be intermediate between the two species (Sauer,
1972). Pembagrass is used in lawns in Kenya (Bogdan, 1970),
Ghana and Uganda (Sauer, 1972), and India (Sundararaj et al.,
1971) and is also a useful pasture grass. The pembagrass USDA
introduction PI-365031 is very coarse textured, and occasionally
the leaf blades are plicate (Busey, 1977, unpublished observations).
Las-aga, S.
micranthum (Desv.) C. E. Hubbard, is a widely distributed
strand pioneer of the Indian Ocean and the South Pacific.
It occurs on open sandy beaches, in the salt spray of coralline
limestone, and other coastal habitats of small islands, but
also extends inland to shaded woodlands and inhabited areas
such as village streets and house yards (Sauer, 1972). In
Guam it is considered an excellent pasture and lawn grass
and is propagated by stolon cuttings (Safford, 1905). Other
than S. secundatum, S. dimidiatum,
and S. micranthum, the four remaining species of Stenotaphrum
are described only from herbarium specimens, not from other
firsthand accounts, and are not cultivated. S. helferi
is distributed from Malaysia through Southeast Asia to southern
China, including Hainan Island. It occurs along forest paths
and serves as good pasture.
The origin of
St. Augustinegrass is unknown. Its distribution has been described
as "part of a larger migrational mystery involving . . . other
cosmopolitan seashore grasses that lack proven capability
of long-range sea dispersal" (Sauer, 1972).
One hypothesis
is that St. Augustinegrass originated in the Old World tropics,
in the center of diversity for the genus, specifically the
coastlines and islands of the Indian Ocean, and that Europeans
later brought it to the New World during the post-Columbian
era. The hypothesis of introduction by Europeans may not explain
the diversity of St. Augustinegrass in the New World, unless
there were multiple early accidental introductions by Europeans.
An alternative
New World origin hypothesis for St. Augustinegrass is consistent
with the early time of the first description of St. Augustinegrass,
1788, from a South Carolina collection (Sauer, 1972), and
even earlier collections in the New World. For example, it
was collected by Dale in the Bahamas, about 1730; by Browne
in Jamaica, about 1750; and by Commerson in Brazil and Uruguay,
in 1767 (Sauer, 1972). St. Augustinegrass has considerable
diversity in cultivated and adventive populations in the West
Indies and southern United States (Busey et al., 1982a). For
example, based on herbarium specimens, by the 1800s St. Augustinegrass
had a wide distribution in North America and showed racial
divergence. The divergence of long-internode plants of the
Longicaudatus Race in Florida, and short-spikelet plants of
the Breviflorus Race in other southeastern states such as
Louisiana (Busey et al., 1982a), suggests a long residence
of St. Augustinegrass in the New World.
A third hypothesis
is that St. Augustinegrass had an Old World origin and was
brought to the New World before the time of European migration,
by an early transoceanic dispersal predating the European
voyages of discovery. This would be consistent with its early
appearance in other distant places; for example, it was collected
by Beauvois in 1787 in Ghana and Nigeria; by Menzies in 1798
from Kauai, Hawaii; and by Cunningham in 1822 in Australia.
Taxonomy and
Geography
Any attempt to
improve St. Augustinegrass genetically would be haphazard
without an understanding of the existing genetic variation,
which is not smooth, but punctuated into clusters of similar
genotypes (Fig. 2). If the clustering were ignored, quantitative
expectations of genetic advance would be biased because assumptions
underlying heritability would be violated. For example, while
quantitative measures of genetic variation assume normal distribution,
under clonal selection of clustered genotypes it is possible
to make rapid initial genetic improvement as the number of
taxonomic groups or ploidy levels is narrowed. But if there
is not sufficient variation within genotype clusters, further
advance may be difficult. In the extreme, attempts at genetic
improvement may be confounded by the occurrence of duplicates
of existing cultivars in the population under selection. Finally,
by providing a natural classification, clustering helps predict
the occurrence of useful alleles (Busey et al., 1982a) for
documented traits such as chinch bug resistance (Busey, 1995a),
disease resistance (Atilano and Busey, 1983), herbicide resistance
(Busey, 1993), nematode resistance (Busey et al., 1993), drought
resistance (Busey, 1996), and shade tolerance (Busey and Davis,
1991).
Morphotype clusters
of St. Augustinegrass (Fig. 2 a-e) have been designated variously
as "Groups", "Races" (Busey et al., 1982a; Busey, 1986), and
"demes" (Sauer, 1972). As an overview to the classification
system, ploidy levels, e.g., 2n=18, are first subdivided into
Races, and Races are subdivided into Groups, which contain
multiple cultivars and breeding populations (Busey et al.,
1982a). Most cultivars are diploids (2n = 18), and diploids
are subdivided into the Breviflorus Race and the Longicaudatus
Race.
The Breviflorus
Race (Busey, 1986) is widely represented among weedy and adventive
populations, and they have high (over 60%) seed set (Busey
and Center, 1979, unpublished data). Within this race, the
Gulf Coast Group (Fig. 2d) is a moderately homogeneous assemblage
of genotypes with green stolons and white stigmata, present
since at least the mid-1800s in the southeastern US, north
of peninsular Florida. The Gulf Coast Group was first clearly
represented in an 1868 collection [ALABAMA: Sandy shores of
Mobile Bay, Point Clear, along the seashore from E. La. to
N. Carol. August 1868, collector Mohr s.n. (AL)]. The Gulf
Coast Group appeared more frequently by the 1890s in Louisiana.
The Gulf Coast Group occurs in protected locations north of
the Piedmont, such as old lawns in Memphis, Tennessee, and
Corinth, Mississippi. These are sources of cold tolerant germplasm
(J. V. Krans, 1984, personal communication). The Gulf Coast
group is endemic to the southeastern United States, and includes
contemporary cultivars 'Texas Common' and 'Raleigh'.
The Dwarf Group
(Fig. 2c), another subcategory of the Breviflorus Race, includes
genotypes with generally strong anthocyanin pigmentation in
the stolons, purple stigmata, and dark green leaf blades (Busey
et al., 1982a). Genotypes of the Dwarf Group generally have
shorter leaves and inflorescences than the Gulf Coast Group.
Artificial introgressants between the Gulf Coast Group and
the Dwarf Group have produced suitable hybrids, some of which
are represented by the shade tolerant cultivars developed
by O.M. Scotts & Sons (Busey and Davis, 1991). One example
is 'Seville' (Riordan et al., 1980), the first St. Augustinegrass
released with a known pedigree, that is, both male and female
parents are known.
Longicaudatus Race
genotypes (2n=18) have elongate stolons (Busey, 1986) and
long leaves (Fig 2b). This race is probably synonymous with
the Natal-Plata deme (Sauer, 1972). Longicaudatus Race genotypes
in older lawns and pastures have been assigned to 'Florida
Common' (Busey et al., 1982a) and include the cultivar 'Roselawn'
(Allen and Kidder, 1971). In Florida, Longicaudatus Race plants
were collected by 1845 in Manatee County, Florida [FLORIDA:
BM: Am Strande, Terraciera Bay, July 1845, Rugel 370 (F,MO,US)],
by 1848 in Key West [FLORIDA: Key West, Herb. Chap. "Prob.
Torrey mis. 1848" (MO)], and by 1894 in central Florida, where
it was regarded as "valuable in pastures" [FLORIDA: St. Augustine
grass. Orlando, Fla., 23 April 1894, Northey 2570 (US)]. This
race grows in remote areas in Everglades National Park, e.g.,
from Highland Beach to East Cape Sable (Busey, et al., 1982),
which was inhabited briefly by Anglo-Americans, around 1900
(Tebeau, 1968). Everglades Experiment Station, University
of Florida, distributed the cultivar Roselawn in 1942 and
1943 (Allen and Kidder, 1971). It has an open habit of growth
and does not form a dense sod (Busey, 1977, unpublished data).
Although not making acceptable lawns, the Longicaudatus Race
apparently has long-term survival ability in low maintenance
habitats.
Polyploidy
Polyploid St. Augustinegrasses
were first identified by Long and Bashaw (1961) who described
sterile triploids (2n=27) with irregular meiosis. They were
designated the Cape deme by Sauer (1972) who identified their
first collection in 1791 at the Cape of Good Hope, and their
use in lawns in the Republic of South Africa by 1900. In fact,
use of polyploids in lawns occurred in Florida by 1892 [FLORIDA:
Cultivated, Leesburg, 6 June 1892, P. H. Rolfs 1008 (US)].
Since 1900, the polyploids have spread most often in association
with intentional introductions and cultivation through vegetative
propagation. 'Bitterblue', a Cape deme genotype (Fig. 2a),
was the foundation for the commercial sod industry in Florida,
starting in the 1920s (Busey and White, 1993). Although Bitterblue
is a sterile clone, slight but detectable genetic variation
exists (Busey, 1986). Another cytologically sterile variant,
Floratam (2n = c. 32, Busey, 1979), was released for its combined
resistance to the St. Augustine Decline Strain of Panicum
Mosaic Virus (PMV-SAD) and the southern chinch bug (Horn et
al., 1973). Floratam St. Augustinegrass (Fig. 2e) became so
popular that by 1980-81, it represented 77% of commercial
sod in southeast Florida and 21% of lawn areas (Busey, 1986).
An unusual 2n=30
polyploid variation was discovered, among African introductions,
with normal bivalent chromosome pairing (Fig. 3) at diakinesis
and normal set seed (Busey, 1990b). From this germplasm, the
cultivar FX-10 was developed with resistance against a virulent,
Floratam-killing race of the southern chinch bug (Busey, 1993).
The simplest cytological origin for the African polyploids
would be allotetraploidy. A 2n=12 progenitor has not been
discovered, and seems unlikely, considering that x=9 or 10
is the basic chromosome number of the Paniceae (Gould, 1968).
Anomalous chromosome counts have been found, however, for
S. dimidiatum: 2n=36 from Sri Lanka (Gould and
Soderstrom, 1974), 2n=48 from Malagasy (Sauer, 1972), 2n =
54 for PI-365031 from the Republic of South Africa (Busey,
1990b), and 2n = c. 60 for FL-2195 from Mauritius (Busey et
al., 1993). It is possible that polyploidy originated in Stenotaphrum
occasionally and by different mechanisms. Polyploidy is important
in the development of other warm-season turfgrasses in addition
to St. Augustinegrass; examples are bahiagrass, Paspalum notatum
Flügge and bermudagrasses, Cynodon spp. (Busey, 1989).
Encouragingly,
taxonomic classifications based on cytology and chemistry
are congruent, suggesting that they are natural. Polyploids
have no detectable activity for uridine diphosphate (UDP)
glucose pyrophosphorylase (Green et al., 1981). Polyploid
genotypes studied included Bitterblue, 'Floralawn' (FA-108),
Floratam, Floratine, FA-118, PI-290888, PI-300127, and PI-300130,
based on direct counts of chromosomes (Busey, 1990b) and/or
racial grouping (Busey, 1986). In contrast, 17 diploid St.
Augustinegrasses have detectable UDP glucose pyrophosphorylase
activity, and so do S. dimidiatum accessions
PI-289729 and PI-365031 (Green et al., 1981). Among diploids
in the latter study, all with low adenosine diphosphate (ADP)
glucose pyrophosphorylase activity were of the Gulf Coast
Group; most with high activity were of the Dwarf Group (Busey
et al., 1982a).
ADAPTIVE POLYMORPHISMS
Physiology
and Environmental Stresses
Polymorphisms
among St. Augustinegrasses have been detected for many physiological
and morphological traits, including isozymes (Green et al.,
1981), leaf extension rate and stomatal density (Atkins et
al., 1991), leaf pubescence (Busey, 1990b), several morphological
and pigmentation traits (Busey et al., 1982a; Busey, 1986),
and lethal temperature and winter survival (Philley, 1994;
Philley et al., 1998). No differences among genotypes have
been observed for mowing energy requirement (Fluck and Busey,
1988).
Adaptive and morphological
variations in St. Augustinegrass are associated with chromosome
differences. The most conspicuous visible differences between
ploidy levels are that diploids have narrower, thinner, more
translucent, brighter green leaf blades, while polyploids
have coarser, thicker leaf blades which are more opaque and
less saturated in color (Busey, 1986, 1993). Compared with
diploids, polyploid leaf blades look grayish blue-green in
lawns. Diploids of the Breviflorus Race have lower growth
habit and more rapid ground covering ability (Busey et al.,
1982a). Their growth habit is more highly branched, which
results in earlier sod maturity, earlier and easier harvest,
but greater risk of thatch problems in the established landscape
(Busey, 1979, unpublished data). In small experimental plots,
such as those in the National Turfgrass Evaluation Program
(NTEP), diploids receive higher turfgrass quality scores than
polyploids, particularly during the first year of evaluation
(Busey, 1985), which can be deceptive for estimating long-term
performance. Polyploids such as Floratam, the main cultivar
in Florida, perform unacceptably for turfgrass quality in
small plots. Most population improvement has been done on
diploids, while polyploid cultivars (e.g., Bitterblue and
Floratam) are often selections or seedlings of unknown paternity
(e.g., Horn et al., 1973).
Shade tolerance
differences exist. Seville, DelMar, and Jade provide superior
quality, compared with Floratam and Floralawn, under 21% relative
irradiance (full sunlight = 100%). Shade was due to a mixed
tree canopy (Busey and Davis, 1991). While photosynthetic
rates among cultivars are similar at 45% or higher relative
irradiance, at 29% relative irradiance, the photosynthetic
rates of Floratam and Floralawn are reduced to less than half
of maximum, which is also less (P<0.05) than Floratine and
Seville, at the same shade level (Peacock and Dudeck, 1993).
Some polyploids, such as Floratam, grow very poorly in the
shade, and genotypic differences in shade adaptation are evident
between 21% and 29% relative irradiance (Busey and Davis,
1991; Peacock and Dudeck, 1993). This could be largely an
expression of leaf height, because polyploids are taller (Busey,
1991, unpublished data).
Compared with
diploids, polyploids are more resistant to drought based on
wilt avoidance due to deeper or more effective root systems,
rather than reduced evapotranspiration. Evapotranspiration
rates in an environmental chamber differ among cultivars,
ranging from 6.7 mm day-1 to 8.1 mm day-1; however, differences
among cultivars are not detected in the field, based on the
average of 3 years (Atkins et al., 1991). Likewise in weighing
field lysimeters, St. Augustinegrasses do not differ in evapotranspiration
(Miller and McCarty, 2001).
Among St. Augustinegrass
genotypes differing in drought survival, extent of wilt is
associated with canopy loss following irrigation suspension
(Busey, 1986). Under conditions of unrestricted rooting in
the field, where there is a water table at 1.45 m, 'FX-10'
has significantly less wilt than Floratam and other cultivars.
When the root systems are confined at 0.75 m, however, the
number of days to wilt for FX-10 was 6.7, which is not significantly
different than Floratam, 6.0 days, but is greater than 'Palmetto,'
4.8 days (Miller and McCarty, 2001).
These results
are consistent with the hypothesis that FX-10 avoids wilt
by deep rooting, provided there is room for deep rooting.
Compared with Floratam, FX-10 was able to maintain a superior
leaf water potential at the first end point (water exudation
from the veins of the cut leaf edge), but not at the second
end point (darkening of the leaf) (Miller and McCarty, 2001).
FX-10 has a prominent, heavily suberized endodermis (Fig.
4), which may be related to root permeability to water.
Floratam, the
only polyploid extensively studied for freezing tolerance,
has no detectable cold acclimation (Fry et al., 1991). Lethal
temperatures for regrowth are -4.5° C and -6.0° C for Floratam
and Raleigh, respectively; electrolyte leakage differences
are similar, but smaller (Maier et al., 1994a, 1994b). These
differences are significant in the field. Winter-kill occurs
to sensitive cultivars such as Floratam following temperatures
of -9° C to -7° C (Busey, 1990a), yet Floratam also has been
reported to survive as low as -15° C (Wilson et al., 1977).
Raleigh St. Augustinegrass, with higher freezing tolerance
than FX-332 or Floratam, is the only cultivar that acclimates
to cold (Maier et al., 1994b). Differential thermal analysis
(DTA) is highly correlated, r = 0.96, with field survival
rating (Philley et al., 1995).
Cultivars differ
in salinity response. For example, Seville is more tolerant
of salinity than Floratam, Floratine, or Floralawn based on
hydroponic culture (Dudeck et al., 1993; Peacock et al., 1993;
Smith et al., 1993) but not based on whole plant microculture
(Smith et al., 1993).
Biotic Stresses
Genotypic differences
occur in resistance to the southern chinch bug (Reinert and
Dudeck, 1974); resistance to the sting nematode, Belonolaimus
longicaudatus (Busey et al., 1993); resistance to the
St. Augustine Decline Strain of Panicum Mosaic Virus (PMV-SAD)
(Horn et al., 1973); infectivity by Sclerophthora macrospora
(Sacc.) Thirum., Shaw, & Naras. (Grisham et al, 1985), the
cause of downy mildew disease; and susceptibility to Pyricularia
grisea (Cke.) Sacc., the cause of gray leaf spot disease
(Atilano and Busey, 1983).
No differences
among genotypes have been observed for resistance to brown
patch disease (Hurd and Grisham, 1983), caused by Rhizoctonia
solani Kuhn; nor resistance to Gaeumannomyces graminis
(Sacc.) Arx & D. Olivier var. graminis, the causal
organism of take-all root rot of St. Augustinegrass (Elliott
et al., 1993); nor resistance to the lance nematode, Hoplolaimus
galeatus (Cobb) Thorne (Henn and Dunn, 1989; Giblin-Davis
et al., 1995). Despite differences in suitability of St. Augustinegrass
genotypes as hosts to the lance nematode, based on nematode
reproduction, even at populations exceeding 10,000 nematodes
g-1 soil, there is no measurable effect of lance nematodes
on roots or shoots.
Much of the variation
in resistance to biotic stresses is accountable by ploidy
level. Compared with diploids, polyploids are more resistant
to the southern chinch bug (Busey, 1990b; Busey and Zaenker,
1992; Reinert et al., 1986) and the sting nematode (Busey
et al., 1993). Polyploids are less preferred by Lepidoptera
than diploids (Busey et al., 1982a). Polyploids of the Bitterblue
Group are highly susceptible to gray leaf spot disease (Atilano
and Busey, 1983). Plant breeders should be encouraged by the
large genotypic variations revealed in germplasm screenings.
Yet when variances in these studies are partitioned into ploidy
levels, and genotypes nested within ploidy levels, often the
vast majority of genetic variation is between ploidy levels
(e.g., Busey and Zaenker, 1992; Busey et al., 1993). This
suggests that some of the detectable genetic variation is
not readily usable unless methods can be developed for gene
exchange between ploidy levels.
Besides locating
sources resistant to major pests, the dynamics of the host-pest
relationship, and the most efficient method of screening need
to be understood. This is illustrated by the southern chinch
bug, an insect with variable populations. Floratam, released
for its chinch bug resistance (Horn et al., 1973), remained
free from economic damage by chinch bugs for 12 years, according
to sod growers and commercial lawn applicators. The resistance
of Floratam was confirmed by repeated laboratory screenings
(reviewed by Quisenberry, 1990). In 1985, however, southern
chinch bugs killed large areas of Floratam. The damaging chinch
bugs were shown to be a population with virulence to Floratam
(Busey and Center, 1987). Introduced African germplasm provided
the foundation for a new cultivar, FX-10, which remains resistant
to different chinch bug populations (Busey, 1990b; Cherry
and Nagata, 1997). Excreta residue deposited on aluminum foil
is a rapid method for assessing chinch bug host suitability
of St. Augustinegrass germplasm (Busey and Zaenker, 1992).
Both excreta residue and oviposition rate have high association
with extent of field damage from natural infestations, r2
= 0.57 and 0.67, respectively (Busey, 1995a).
Pathogens also
vary in virulence, which may explain differences in disease
incidence in different regions. St. Augustine decline isolates
of Panicum Mosaic Virus (PMV-SAD) vary serologically, which
may explain variable lethality to St. Augustinegrass lawns
(Holcomb et al., 1989). Isolates may also represent mixtures
of strains or serotypes, making resistance screening more
unpredictable. Resistance screening based on a single isolate
may be a poor representation of pathogen variation, and lead
to inaccurate estimates of host susceptibility.
INTRODUCTION,
SELECTION, AND BREEDING
Germ Plasm
Resources
In 2001, 23 foreign
introductions of Stenotaphrum were available for breeders
in the National Plant Germplasm System (USDA, ARS, National
Plant Germplasm Program, 2001). These clonal plants included
two accessions (PI-289729 and PI-365301) of S. dimidiatum
(incorrectly labeled S. secundatum), and the
rest S. secundatum. The most recently introduced
genotypes available for distribution were two collected by
Dr. Milt Engelke from China, added in 1993; the next most
recently added genotype was in 1979. In addition, four plants
submitted by Mr. Tobey Wagner, and two plants from Dr. Jeffrey
V. Krans, are awaiting release from quarantine. Much of the
potential germplasm of St. Augustinegrass occurs in pastures,
especially in coastal Africa, from Kenya to the Cape of Good
Hope (Chippindall and Crook, 1976), the West Indies (Busey
et al., 1982a), and Oceania (Sauer, 1972).
Released cultivars
and active breeding populations, outside the minuscule US
collection, represent most of the germplasm available to breeders.
Most of the S. secundatum genotypes used in
breeding programs represent the Dwarf Group, with little attention
to the African polyploids (Busey et al., 1982a). It is not
known what other groups lay undiscovered. For the Breviflorus
Race, which is so extensively used in breeding programs, genetic
variation is readily available in adventive populations in
the southeastern United States (Busey et al., 1982a).
St. Augustinegrass
has been naturalized since at least the 1700s in North and
South America, Africa, and the Pacific, and exhibits considerable
phenotypic variation throughout its range. Because of this
antiquity, there is probably no area where there is not some
useful genetic variation. Even far outside the presumptive
center of origin in the Indian Ocean area, germplasm collections
of St. Augustinegrass may be very useful, because they may
represent relict types that no longer occur in the natural
range. A caution in germplasm collections of St. Augustinegrass
is to be diligent to cull out duplicates that represent widely
distributed clonal cultivars. Also, because St. Augustinegrass
is propagated and marketed in an active, vegetative condition,
breeders and germplasm managers must also be aware of the
perils of accidentally dispersing systemic and attached disease
organisms, such Sclerophthora macrospora, Gaeumannomyces graminis
var. graminis, as well as St. Augustine Decline Strain of
Panicum Mosaic Virus (PMV-SAD), and the sting nematode.
Pembagrass, S.
dimidiatum, PI-365031 has resistance to gray leaf spot
disease caused by Pyricularia grisea (Cke.) Sacc. (Atilano
and Busey, 1983) and the southern chinch bug (Busey, 1990b);
S. dimidiatum FL-2195 has resistance to the
sting nematode (Busey et al., 1993). Therefore S. dimidiatum
is a good first candidate for wide crosses and other methods
for gene transfer.
Breeding and
Selection Techniques
St. Augustinegrass
is easy to hybridize artificially (Philley et al., 1993).
Inflorescences are photoperiod-controlled (Dudeck, 1974),
and flowering occurs first in the center of the inflorescence,
and progresses predictably in both directions. Anthesis in
most genotypes occurs soon after sunrise, but anthesis of
S. dimidiatum is at night. At the University
of Florida, Fort Lauderdale, parchment pollinating bags were
placed over inflorescences one day before anthesis, with the
plants generally in containers in a greenhouse, although bagging
of plants in field plots was also performed. The relatively
large spikelets were easily emasculated with a pair of forceps,
which was done in the morning as the anthers emerged. By also
removing unused spikelets, and unused portions of the inflorescence,
it was easier to keep track of crosses, and less likely to
have stray pollen in a bag. Any spikelets with already dehisced
anthers were removed from the inflorescences. Crosses were
made using pollen transferred to the hand-emasculated florets.
Pollinated spikelets were marked with an indelible marker.
In addition, spikelet positions were numbered and recorded,
so that a record of shriveled stigmata (an indication of effective
crossing) could later be associated with individual seeds
harvested. The reaction of the stigmata was recorded one day
after pollination, and the bags removed.
Seed is set and
easily produced within ploidy levels. Bivalent-pairing polyploids
(2n = 30) from southern Africa produce 43% to 70% seed set
(Busey, 1990b). Diploids (2n = 18) of the Breviflorus Race
produce over 60% seed set. However, ploidy level differences
impede the full use of germplasm; intended crosses between
different ploidy levels have not been successful (Busey, 1981,
unpublished data). The most successful St. Augustinegrass
in Florida, Floratam, normally produces no seed. However,
in 1983 seed were obtained from Floratam growing in a greenhouse,
open-pollinated by 2n = 30 African parents. Among the progeny,
several had laminar hairs similar to the putative male parents.
One of the Floratam progeny, FX-5, had reduced oviposition
by the Polyploid Damaging Population (PDP) southern chinch
bug (Busey, 1990b), evidence for chinch bug resistance conferred
by the African male parents.
Because antibiotic
resistance to the southern chinch bug has not been discovered
among diploids (Busey and Zaenker, 1992; Reinert et al., 1986),
embryo rescue or protoplast fusion might be used to transfer
this trait across the ploidy barrier. The caryopses of St.
Augustinegrass mature more quickly than Zea mays L. At 9 days
after pollination, the St. Augustinegrass embryo (Fig. 5)
has developed leaf primordia and vasculature, and is nearly
half of its mature length (2.05 mm). By 10 days after pollination,
radicle development has begun. In contrast, in Z. Mays the
leaf primordia form at 12 days, while in Eragrostis curvula
this occurs at 5 days.
Somatic mutations
are easily produced in St. Augustinegrass sprigs using gamma
rays, and 3000 to 4500 rads is the appropriate dosage to generate
high mutation rates and adequate sprig survival, depending
on the cultivar (Busey, 1980). Complete plant regeneration
has been accomplished for St. Augustinegrass from callus (Kuo
and Smith, 1993).
The biggest challenge
in breeding St. Augustinegrass is that it is a perennial,
and evaluation is difficult. Field evaluation must be long-term,
exposing genotypes to a range of chronic natural problems
(e.g., nematodes and thatch) and acute environmental and biotic
problems (injury from drought and chinch bugs). St. Augustinegrass
does not exhibit some pest problems, such as sting nematode,
for at least two years after establishment (Busey et al.,
1991), and southern chinch bug infestation typically begins
in susceptible cultivars about 1.5 years after plug planting.
Attempts to accelerate the progress of evaluation by prescreening
for plant characteristics in containers was not successful,
as no correlation was found between container performance
and field performance (Busey, 1981, unpublished data).
Inheritance
Diploid (2n =
18) St. Augustinegrass has normal paired-factor inheritance,
based on Mendelian 3:1 ratios for stigma color observed in
segregating progeny, consistent with an hypothesis that purple
stigma is dominant to white (translucent) (Table 1). A white
stigma irradiation-induced mutation was derived from a heterozygous
purple-stigma genotype (Busey, 1980), which supports simple,
diploid inheritance control.
Variegation is
simply inherited. For example, the selfed progeny of normal
green-leafed plant FA-243-39 were 7 variegated and 20 normal,
consistent with an hypothesis that variegation is a single
recessive, giving an expected 1:3 ratio. However, a second
gene may also be involved, because the selfed progeny of normal
green-leafed plant 365032-8F231 were 38 variegated and 42
normal, which is consistent with the variegated trait being
recessive on two epistatic loci, giving an expected 7:9 ratio.
Variegated St. Augustinegrass, which has invalidly been referred
to in horticultural encyclopedias as Stenotaphrum variegatum,
was documented by the famous agrostologist Dr. Agnes Chase
from a hanging pot in a greenhouse in Garfield Park, Chicago
[ILLINOIS: Chicago. 27 October 1915; Chase, s.n. (USNAT)].
Variegated St. Augustinegrass was used as a model species
for studying chloroplast enzymes (Suzuki et al., 1986). The
variegated mutation has appeared independently in different
germplasms, for example, in turf exposed to oxidizers such
as laundry detergent and swimming pool water (Busey, 1980,
unpublished observations). Other genetic traits are not well
understood.
Reproduction
Deliberate propagation
of St. Augustinegrass is usually vegetative, by stolon cuttings,
plugs, and sod. The only commercially available cultivars
are thus clones. Grown as a monoculture, St. Augustinegrass
remains vulnerable to pest evolution (Busey and Center, 1987).
Efforts to develop seeded cultivars might enhance genetic
diversity, but have not been successful, despite repeated
attempts. For example, in 1974, Curran L. Garrett received
a plant patent for a heavily seed-producing St. Augustinegrass.
Also, in the early 1990s Pennington Seed marketed seed from
St. Augustinegrass, calling it 'Raleigh-S.' Unfortunately,
genotypes that are prolific seed producers are often esthetically
unacceptable in regions with an extended growing season (Busey,
1984, unpublished data). In addition, inbreeding depression
occurs in St. Augustinegrass, and the seed produced from a
clonal monoculture must, by nature, be inbred. Even ignoring
the genetic problems of seed production in St. Augustinegrass,
seed yield is low and it is very difficult to remove the caryopses
from the corky rachis segments. An alternative to seeded cultivars
would be clonal blends of cultivars differing in host resistance.
An esthetically compatible blend might be protected from pest
dispersal and outbreak, either directly because of the dilution
of host density, or indirectly because the natural selection
pest virulence would be delayed by the genetically heterogeneous
host.
History of Breeding
and Population Improvement
Organized breeding
of St. Augustinegrass has occurred on few occasions. This
is accountable in part because it is primarily a lawn grass,
and not important for golf or sports turf, thus sources of
research funds have been minimal. For example, between 1983
and 1997 the United States Golf Association (USGA) funded
$3.86 million for turfgrass breeding of bermudagrass, Cynodon
spp.; zoysiagrasses, Zoysia spp.; seashore paspalum,
Paspalum vaginatum Swartz; buffalograss, Buchloë
dactyloides (Nutt.) Engelm.; And creeping bentgrass, Agrostis
palustris Huds. No funds were allocated, nor proposals
solicited, for St. Augustinegrass improvement.
Commercial breeding
development of St. Augustinegrass has also been limited because
it is a clonal crop, which makes it harder to define and control
the pathway to an effectively large market. St. Augustinegrass
is produced on many independent sod farms. To recoup the cost
of developing intellectual property in St. Augustinegrass,
as well as marketing and quality control, requires effective
licensing to many companies who are in competition with one
another. Potential licensees may vary in size, experience,
and production techniques (e.g., plug production versus sod)
which makes it difficult to standardize licensing requirements
and royalty basis. In contrast, for cultivars of seed-propagated
turf species, such as a perennial ryegrass, Lolium perenne
L., the developer can more readily control the stages of distribution
by concentrating regulation on the more centralized seed production
area, e.g., by subcontracting to growers who sell back to
the developer, who then sells to consumers or brokers. A seed
propagated species has two other advantages in intellectual
property rights. Quality control can be more readily assured
in a seed propagated species because there is a storage period
for quality assessment. Quality control is more difficult
in vegetatively propagated species such as St. Augustinegrass
where the product is perishable and can vary in weed content
and other characteristics during the time it takes to assess
quality. Also, developers of some seed propagated turfgrasses,
such as overseeded perennial ryegrass, expect a lucrative
recurring market from users with annual budgets such as golf
courses, whereas developers of vegetatively propagated turfgrasses
do not expect frequent repurchases.
The Scotts Company
has conducted the major commercial breeding development of
St. Augustinegrass. In research at Scotts farm in Apopka,
Florida, Dr. Terrence Riordan developed numerous clones, several
of which were patented, and three were registered ('DelMar',
'Jade', and Seville). Mr. Tobey Wagner of Sod Solutions (South
Carolina) patented Palmetto St. Augustinegrass, a clonal collection.
A total of 18 plant patents for St. Augustinegrass have been
awarded.
Efforts by public
scientists have involved discovery of clonal types such as
'Floratine' and Raleigh, and discovery of seedlings of partially
unknown pedigree, for example Floratam and 'Floralawn' St.
Augustinegrasses. The author at the University of Florida-Fort
Lauderdale did the only large-scale population improvement,
from 1977 until 1996, when the program was assigned to Dr.
Russell Nagata at the University of Florida-Belle Glade. The
main basis for organized breeding of St. Augustinegrass at
the University of Florida-Fort Lauderdale was a composite
cross population.
From 1978 through
1982, an average of 28 parents per generation (Table 2) were
hybridized randomly to produce offspring populations that
were evaluated in the field in comparison with cultivar standards,
Bitterblue, Floratam, Roselawn, and Seville. The turfgrass
quality mean of cultivar standards was a constant reference
to compare population changes due to composite crossing, selfing,
recurrent selection involving the selection of elite parents,
and vegetative repropagation of plants that had performed
well in earlier trials.
Initial parents
had been chosen to represent taxonomic groups classified from
a worldwide population (Busey et al., 1982a). Parents of each
succeeding generation were chosen based on phenotypic dissimilarity.
In addition to four generations of composite crossing (C1,
C2, C3, and C4), several selfed populations were also created
(e.g., S1, C1S1, etc.). Recurrent selection populations were
created (R1 and R2) from elite parents that were chosen based
on individual plant performance or progeny performance, and
vegetatively repropagated populations (V1, V2, and V3) were
chosen for reevaluation based on their prior superior performance.
On several dates
during the first 14 months after field planting, plots were
evaluated for turf quality, a combination of adaptive and
esthetic traits, with 10=complete coverage, deepest leaf color,
and most dense, low, uniform habit; 7=acceptable coverage,
color, and habit; and 1=plant dead. Because some populations
were evaluated with only a single plot per genotype, and other
populations were evaluated in randomized complete blocks,
the comparison of populations to the mean of cultivars was
on the basis of population individual plot values, rather
than population genotype means. Cultivar standards were, however,
replicated.
Composite crosses
had 20% of plots with turf quality ratings exceeding the mean
of cultivars, which was almost the same fraction as the initial
parents, 17% (Table 2). Genotypic variances did not change
under composite crossing in the absence of selection. For
example, C3 (which was evaluated in three replicates) had
a genotypic variance for turf quality of 1.30, compared with
1.43 for the P1 parents. Selfed populations were inferior,
as would be expected for a normally cross-pollinated species;
only 2% of plots exceeded the mean of four cultivars. In related
work, gray leaf spot disease severity was higher for an open-pollinated
and probably inbred offspring of a Gulf Coast accession than
for the parent (Atilano and Busey, 1983), confirming the problems
of inbreeding St. Augustinegrass.
Narrow-sense heritability
for turf quality was estimated from midparent-offspring regression
and was significant (P < 0.05) in two cases, C2 regressed
on C1 (0.44) and C3 on C2 (0.66), but was not significant
in two cases (C1 on P1 and C4 on C3). Recurrent selection
based on crosses of elite parents was successful, as the R1
and R2 populations had a high proportion (34%) of plots superior
to the mean of cultivars.
The broad-sense
heritability for turf quality in 60 randomly selected, retested
C3 clones was 0.45 (single-plot basis). The average broad-sense
heritability of hybrids within single replicated experiments
was 0.62. With such high heritabilities, little benefit would
be obtained by replicating in first-stage clonal evaluations.
In support of this conclusion, plots of vegetative selections
that were reevaluated (V1, V2, and V3), and which had been
chosen in most cases from no more than two replicates, were
superior 67% of the time compared with the mean of cultivars.
By not replicating in first-stage evaluations, a larger germplasm
can be screened and subjected to more intensive selection,
even though heritability based on unreplicated selection is
less than heritability based on genotype means.
Composite crossing
was also successful in preserving genetic variation, because
after recurrent selection and after vegetative selection,
adequate genotypic variance was found compared with the original
parents. Genotypic variances for the recurrent populations
R1 and R2 were 1.26 and 1.52 units, respectively, and for
the vegetative selections V1, V2, and V3 were 1.34, 1.22,
and 0.85, respectively.
Other work on heritability
under selection, based on the analysis of a diallel cross,
resulted in estimated narrow sense heritability for lethal
temperature of 0.58, and a range from 0.70 to 0.98 for winter
survival. The two traits are correlated with one another (Philley
et al., 1998). Specific combining ability was generally not
significant.
CONCLUSIONS
Scientific attention
to St. Augustinegrass has been sporadic. In the haste to get
new cultivars to market, basic information such as pedigree
and usable description have not been reported, if they are
even known. Meanwhile, other cultivars have undergone unnecessarily
lengthy test periods prior to release, e.g., 26 years in the
case of Floralawn (Dudeck et al., 1986). The review process
for scientific manuscripts and plant patent applications puts
high emphasis on demonstrating cultivar differences, but a
process for evaluating the applicability of the results to
the field is not available. Repeatedly, field performance
variation is poorly predicted based on laboratory evaluation,
e.g. evapotranspiration, shade tolerance, and turfgrass quality.
Even the process of evaluating St. Augustinegrass cultivars
in tests including the National St. Augustinegrass Test by
the National Turfgrass Evaluation Program (NTEP) has resulted
in systematic biases against coarse-textured cultivars such
as Floratam. Floratam is the best adapted and most widely
used St. Augustinegrass in Florida, even though it receives
poor turfgrass quality ratings in most field trials.
In other instances,
e.g., differential thermal analysis for assessing freezing
resistance and excreta residue for assessing host suitability
to the southern chinch bug, laboratory criteria have high
correlation with field traits, and are more efficient for
screening than waiting for natural stresses to occur. Scientists
have developed screening techniques for traits that are relatively
easy to assess, while one of the most difficult adaptive problems
in turf, i.e., shade, is infrequently studied (Fig. 6). Most
turf evaluation environments are in full direct sun. In the
absence of accurate scientific information, marketers of proprietary
St. Augustinegrass cultivars normally make the same claims
of superiority, for drought resistance, shade resistance,
and chinch bug resistance, for all new cultivars. At the least,
landscape plantings of specific cultivars should be revisited
a few years after establishment, to determine actual performance
based on the original expectations.
Finally, there
is the problem of limited funding of research for lawn grasses.
Limited funds are available from state and governmental agencies
to do targeted work on special problems, such as water conservation
research funded by various water authorities. Such agencies
have been ambivalent in recognizing the importance of turf
in the environment, and often seek to replace turf with groundcovers,
rather than to distribute and promote useful irrigation technology.
In other cases, proprietary interests have contracted for
limited research on specific traits of interest in preparing
patents and marketing of found cultivars, but they have not
funded the breeding development of new genotypes. Meanwhile,
the State Agricultural Experiment Stations, which are responsible
for the development of publicly released cultivars, have in
some cases failed to submit successful cultivars for evaluation
in the National Turfgrass Evaluation Program, and in other
cases have failed to maintain the original Breeder's Stock
of released cultivars. A public commitment is needed to the
study of St. Augustinegrass, as a versatile plant that provides
the primary green landscapes and erosion control for tens
of millions of people.
Acknowledgement
This research was
supported by the Florida Agricultural Experiment Station,
and approved for publication as Journal Series No. R-08303.
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Fig. 1. Leaf blade
transverse section of 'Roselawn' St. Augustinegrass showing
the dense Krantz bundle sheath cells surrounding each vascular
bundle, an indication of the C4 photosynthetic pathway.
Fig. 2. Races
and Groups of St. Augustinegrass (Busey et al., 1982a; Busey,
1986). (a) Bitterblue Group, 'Bitterblue'; (b) Longicaudatus
Race, 'Roselawn'; (c) Breviflorus Race Dwarf Group, FA-243;
(d) Breviflorus Race Gulf Coast Group, FL-1933; (e) Floratam
Group, 'Floratam.'
Fig. 3. Pollen
mother cells of St. Augustinegrasses showing entirely bivalent
pairing in diploids (2n=18) and polyploids (2n=30). (a) FX-261
diakinesis (2n=18); (b) FL-1759 diakinesis (2n=30); (c) FA-243
diakinesis (2n=18); and (d) FX-10 metaphase (2n=30).
Fig. 4. Root transverse
section of St. Augustinegrass FX-10 showing the stele with
five xylem elements, surrounded by a densely suberized ring
of endodermis. The cortex cells are partially collapsed. Phloem
cells are small and difficult to discern.
Fig. 5. Seed development
of St. Augustinegrass, Stenotaphrum secundatum,
Scotts-1081, showing longitudinal sections at 7 to 18 days
after pollination (Busey and Center, 1983, unpublished data).
Fig. 6. St. Augustinegrass
in the landscape, FL-1997-6 developed by the writer, forming
a dense turf under the shade of Ficus spp. trees and grapefruit,
Citrus paradisi Macf. at the residence the late Paul Frank,
Golf Course Superintendent, Wilderness Country Club, Naples,
Florida.
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