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Effects of a warmer climate on seed germination in the subarctic

In a future warmer subarctic climate, the soil temperatures experienced by dispersed seeds are likely to increase during summer but may decrease during winter due to expected changes in snow depth, duration and quality. Because little is known about the dormancy-breaking and germination requirements of subarctic species, how warming may influence the timing and level of germination in these species was examined.


Under controlled conditions, how colder winter and warmer summer soil temperatures influenced germination was tested in 23 subarctic species. The cold stratification and warm incubation temperatures were derived from real soil temperature measurements in subarctic tundra and the temperatures were gradually changed over time to simulate different months of the year.

Key Results

Moderate summer warming (+2·5 °C) substantially accelerated germination in all but four species but did not affect germination percentages. Optimum germination temperatures (20/10°C) further decreased germination time and increased germination percentages in three species. Colder winter soil temperatures delayed the germination in ten species and decreased the germination percentage in four species, whereas the opposite was found in Silene acaulis. In most species, the combined effect of a reduced snow cover and summer warming resulted in earlier germination and thus a longer first growing season, which improves the chance of seedling survival. In particular the recruitment of (dwarf) shrubs (Vaccinium myrtillus, V. vitis-idaea, Betula nana), trees (Alnus incana, Betula pubescens) and grasses (Calamagrostis lapponica, C. purpurea) is likely to benefit from a warmer subarctic climate.


Seedling establishment is expected to improve in a future warmer subarctic climate, mainly by considerably earlier germination. The magnitudes of the responses are species-specific, which should be taken into account when modelling population growth and migration of subarctic species.


In Arctic and subarctic regions, which are expected to be affected more by anthropogenic climate change than other regions in the world (ACIA, 2004), the effects of climate warming on germination have hardly been studied (but see Wookey et al., 1995; Molau and Shaver, 1997; Graae et al., 2008). However, the timing and level of germination strongly affect a plant’s recruitment success and may consequently have implications for species migration. Therefore, research on how a warmer climate affects germination is important for our ability to predict population dynamics and future distributions of subarctic species (Higgins et al., 2003; Neilson et al., 2005; Pearson, 2006; Thuiller et al., 2008).

In northern ecosystems, climate warming is not only expected to result in significant increases in temperature, but also changes in the timing and reduction in the depth and duration of the snow cover are projected to occur (ACIA, 2004; Phoenix and Lee, 2004; Keller et al., 2005; IPCC, 2007). The extent of snow cover over Arctic land areas has declined by about 10 % over the past 30 years and model projections suggest that it will decrease an additional 10–20 % before the end of the century (ACIA, 2004). Snow is an excellent insulator (Sturm et al., 2005); therefore a warmer climate is likely to result in colder soils and deeper soil frost during the winter when the soil is not insulated by snow (Groffman et al., 2001; Venäläinen et al., 2001; Stieglitz et al., 2003; Öquist and Laudon, 2008). Moreover, important snow quality changes are projected; for instance, the development of hard packed snow due to changing wind patterns and ice layer formation, due to an increase in thawing and freezing in winter (ACIA, 2004), resulting in less insulation and thus colder soils (Körner, 2003). This may have important implications for recruitment from seed, because in subarctic environments, where there is hardly any current season (pre-winter) germination (Körner, 2003), winter temperatures possibly influence seed germination (Vleeshouwers et al., 1995; Baskin and Baskin, 1998).

In cold climate species, experiencing winter (cold stratification) usually increases germination percentages (Probert, 2000; Körner, 2003; Giménez-Benavides et al., 2005) and often contributes to reducing the temperature needed for later germination (Reynolds, 1984; Densmore, 1997; Shimono and Kudo, 2005). Cold stratification studies of Arctic and alpine species have, however, mainly focused on the duration of the stratification period (e.g. Baskin et al., 2000; Cavieres and Arroyo, 2000; Schütz, 2002) rather than on the stratification temperature. This is usually selected close to 5 °C because this is supposed to be optimal for dormancy-breaking in many species (Bewley and Black, 1994). Some recent studies have used more realistic stratification temperatures close to 0 °C for cold climate species (e.g. Baskin et al., 2000; Schütz, 2002; Shimono and Kudo, 2005; Graae et al., 2008), but this only reflects situations where a thick snow cover provides insulation. The effect of colder winter soil temperatures, reached in the absence of snow, has to our knowledge only been tested in two experiments with stratification under field conditions (Baskin et al., 2002; Graae et al., 2008). However, these included only two species, Empetrum hermaphroditum and Vaccinium uliginosum, and no effects of stratification temperature were observed.

The germination temperature in Arctic and alpine plants is relatively high in comparison with ambient temperatures (Cavieres and Arroyo, 2000; Körner, 2003), which has been thought to be an adaptation to avoid germination in early spring or autumn when the probability of frost is high (Billings and Mooney, 1968; Cavieres and Arroyo, 2000). Further, germination percentages are often positively correlated with the temperature under which seeds are incubated (Baskin and Baskin, 1998; Graae et al., 2008). However, apart from these general trends, detailed knowledge on the germination requirements of (sub)arctic species is lacking (Baskin and Baskin, 1998). This is mainly because most studies use optimal incubation temperatures for testing germinability, often 20/10 °C or 25/15 °C (for a review, see Baskin and Baskin; 1998), independent of the species’ habitat. The use of more realistic temperatures adapted to the environment of the species or the application of gradually changing temperature regimes simulating each month of the growing season is still scarce (but see Baskin et al., 1995; Thompson and Naeem, 1996), although essential to apply germination data from laboratory studies in an ecological context. Moreover, the importance of the timing of germination has often been overlooked, despite its significance for plants from harsh environments where the growing season is highly limited (Schütz, 2002; Deines et al., 2007).

To improve our knowledge of how climate warming may affect germination in subarctic ecosystems, an experiment was set up in which the effects of two realistic cold stratification treatments and three warm incubation treatments were tested. The temperature regimes were derived from real soil temperature measurements in subarctic tundra and were gradually changed over time to simulate different months of the year. As stratification temperatures, winter soil temperatures were used from snow-covered and exposed habitats, the latter to simulate a reduced snow cover in a future warmer climate. The incubation temperatures corresponded to (a) current spring and summer temperatures in subartic tundra, (b) temperatures that are 2·5 °C higher than that, and (c) optimum germination temperatures (20/10 °C). To unravel general patterns, seeds from 20 subarctic species belonging to different functional types were used. Also three boreal tree species were added to test if their current expansion is limited by unsuccessful germination under present subarctic temperatures.

Specific research questions were: (a) How are germination percentage and germination time affected by colder winter soil temperatures, occurring under a reduced snow cover? (b) Does a higher germination temperature (+2·5 °C) affect germination of subarctic species, and if so, how? (c) Are the effects general or species specific? (d) Is the range expansion of boreal tree species currently limited by too-low germination temperatures in subarctic tundra?


Study species

Seeds of 20 species were collected in the surroundings of the Abisko Scientific Research Station (68°21′N, 18°49′E) in subarctic northern Sweden (Table  1 ). The species were selected to belong to different functional types and were important components of the plant communities in the study area. Additionally, seeds of Alnus incana and Betula pubescens, tree species with a boreal distribution, were collected close to Umeå (63°50′N, 20°20′E) and seeds of Pinus sylvestris were ordered from a seed company and originated from Karesuando (68°20′N, 21°53′E). All seeds (except from P. sylvestris) were collected between 15 August and 15 September 2007 and stored dry at room temperature until sowing (6 November 2007). Seeds of fleshy fruits were extracted from the fruits immediately after collection. The seed mass of each species, without any attachments, was determined by weighing four times 30 air-dry seeds (ten seeds for Pinus sylvestris and Vicia cracca because of their large seed size).

Table 1.

Average seed mass and functional type of the species studied

Species Seed mass (mg) Functional type
Alnus incana (L.) Moench* 0·547 Tree
Astragalus frigidus (L.) A. Gray 2·920 Forb (legume)
Betula nana L. 0·223 Shrub
Betula pubescens Ehrh.* 0·164 Tree
Betula pubescens ssp. czerepanovii (Orlova) Hämet-Ahti 0·236 Tree
Calamagrostis lapponica (Wahlenb.) Hartm. 0·328 Grass
Calamagrostis purpurea (Trin.) Trin. 0·092 Grass
Carex rostrata Stokes † 1·319 Sedge
Carex saxatilis L. † 0·702 Sedge
Deschampsia flexuosa (L.) Trin. 0·498 Grass
Dryas octopetala L. 0·335 Dwarf shrub
Empetrum hermaphroditum Lange ex Hagerup † 1·230 Dwarf shrub
Epilobium angustifolium L. 0·065 Forb
Festuca ovina L. 0·245 Grass
Pinus sylvestris L.* 4·600 Tree
Salix glauca L. 0·156 Shrub
Silene acaulis (L.) Jacq. 0·402 Forb
Silene dioica (L.) Clairv. 0·831 Forb
Solidago virgaurea Praecox 0·327 Forb
Vaccinium myrtillus L. 0·214 Dwarf shrub
Vaccinium uliginosum L. 0·218 Dwarf shrub
Vaccinium vitis-idaea L. 0·202 Dwarf shrub
Vicia cracca L. 12·580 Forb (legume)

* Species with a boreal distribution

† Species excluded from further analyses because of very poor germination (<5 %)

Cold stratification and warm incubation treatments

The seeds were subjected to two contrasting cold stratification treatments for 20 weeks. The stratification temperatures (Table  2A ) were based on winter soil temperature measurements (3 cm depth) during 2006 and 2007 in subarctic tundra sites with either a thick insulating snow cover throughout winter (meadow vegetation) or a strongly reduced snow cover (wind exposed poor dwarf shrub heath). The measurements were done approx. 8 km from the Abisko Scientific Research Station at 850 m a.s.l. (B. J. Graae, unpubl. res.). Cold stratification occurred in complete darkness.

Table 2A.

Temperatures during the different weeks of the stratification treatments

Stratification temperature (°C)*
Week of the experiment Equivalent time of year Thin snow cover Thick snow cover
1–4 October–November 0·5 0·5
5–8 December −5 0·5
9–12 January–February −10 0·5
13–16 March to 15 April −5 0·5
17–20 15 April to 15 May 0·5 0·5

* The ‘thin snow cover’ temperatures are based on soil temperature measurements (3 cm depth) during the indicated time periods in wind-exposed poor heath vegetation in subarctic tundra near Abisko (northern Sweden) and the ‘thick snow cover’ temperatures on measurements in meadow vegetation at the same location.

After stratification, the seeds were allowed to germinate under three different incubation treatments: optimum germination temperatures (20/10 °C) (Baskin and Baskin, 1998), current spring and summer soil temperatures in subarctic tundra (control), and temperatures that are 2·5 °C warmer than that (+2·5 °C; Table  2B ). The temperature increase of 2·5 °C is in accordance with the projected temperature rise in Arctic regions (60–90°N) by the year 2050 relative to 1981–2000 (ACIA, 2004). The control incubation temperatures were derived from soil temperature measurements during the spring and summer of 2006 and 2007 in dwarf shrub heath, the dominant vegetation type at the earlier-mentioned location. To simulate the subarctic summer, seeds were kept in 24 h of daylight during incubation, but light intensities were alternated every 12 h coinciding with the temperature intervals. During ‘night’, light was provided with a photosynthetic photon flux density (PPFD) of 25 µmol m −2 s −1 (400–700 nm), whereas during daytime a PPFD of 45 or 110 µmol m −2 s −1 was provided every other day, to simulate overcast and clear days. In the ‘optimum’ treatment, the daytime PPFD was always 110 µmol m −2 s −1 .

Table 2B.

Temperatures during the different weeks of the incubation treatments

Incubation temperature (°C)
Week of the experiment Equivalent time of year Control* +2·5 °C Optimum
21–22 15–31 May 4/0·5 6·5/3 20/10
23–26 June 10·5/2 13/4·5 20/10
27–33 July–August 12·5/4·5 15/7 20/10

* The ‘control’ temperatures are based on soil temperature measurements in dwarf shrub heath vegetation in subarctic tundra near Abisko. Temperatures were alternated every 12 h

For each combination of stratification and incubation treatment, four replications of 30 seeds (ten seeds for P. sylvestris and V. cracca) were sown on moist commercial pot soil in 90-mm-diameter Petri dishes on 6 November 2007. After sowing, the dishes were wrapped with parafilm to reduce loss of water and they were subsequently put in incubators for the cold stratification treatments.

Germination recording

During incubation, the seeds were checked weekly for germination (protrusion of the radicle) and the germinated seeds were removed to reduce counting time in subsequent weeks. After 10 weeks, most species had completed germination but one final germination counting was done after 13 weeks of incubation. The final germination percentages were arcsine transformed to improve normality and stabilize variances. Species with <5 % germination (Carex rostrata, C. saxatilis and Empetrum hermaphroditum) were excluded from further analyses (Table  1 ).

In addition to percentage germination, the mean germination time (MGT) for each of the species was determined in order to distinguish fast from slowly germinating species and to determine how germination speed was affected by the treatments. It was calculated as

where ni is the number of seeds that germinated within consecutive intervals of time, ti the time between the beginning of the test and the end of a particular interval of measurement, and N the total number of seeds that germinated (Deines et al., 2007).


Percentage germination

Significant differences in germination percentages were observed between the functional types (ANOVA, F5,474 = 16·0, P < 0·001), with the poorest germination in the grasses (20 %) and the highest germination in the forbs (56 %; Fig.  1 A). Germination percentage slightly increased with seed mass (linear regression, F1,478 = 30·83, P < 0·001, R 2 = 0·06).

(A) Germination percentages and (B) mean germination times for the different functional types. Data are averages over all treatments (n = 24). Bars denote ± s.e. Different letters indicate significant differences between functional types at the P < 0·05 level (Tukey multiple comparison test).

Germination percentages were significantly affected by the incubation and stratification treatments. In general (all species combined), the ‘thick snow cover’ stratification treatment resulted in more germination (43 %) than the ‘thin snow cover’ treatment (39 %; Fig.  2 A and Table  3 ). In addition, germination percentages were higher in the ‘optimum’ (45 %) incubation treatment than in the ‘ + 2·5 °C’ (39 %) and ‘control’ (39 %) treatments (no difference between +2·5 °C and control; Fig.  2 A). The lack of a significant interaction effect between incubation and stratification (Table  3 ) suggests that the effects of both are additive, which is supported by the highest germination percentage when ‘thick snow cover’ stratification was followed by ‘optimum’ incubation (Fig.  2 A).

Table 3.

Results of two three-way ANOVAs on the effects of incubation treatment, stratification treatment and species identity on germination percentage and on mean germination time

Germination percentage Mean germination time
Factor d.f. F-value P-value d.f. F-value P-value
Incubation (I) 2, 360 12·80 < 0·001 2, 333 407·67 < 0·001
Stratification (S) 1, 360 9·56 0·002 1, 333 11·93 0·001
Species 19, 360 95·19 < 0·001 19, 333 60·96 < 0·001
I × S 2, 360 2·15 0·118 2, 333 0·43 0·653
I × species 38, 360 4·30 < 0·001 38, 333 3·42 < 0·001
S × species 19, 360 4·43 < 0·001 19, 333 1·45 0·103
I × S × species 38, 360 1·18 0·218 38, 333 0·50 0·994

Significant effects (P < 0·05) are in bold.

(A) Germination percentages and (B) mean germination times for the different cold stratification and incubation treatments. See Table 2 for the incubation and stratification temperatures. Data are averages over all species (n = 80). Bars denote ± s.e. Different letters indicate significant differences between incubation treatments at the P < 0·05 level (Tukey multiple comparison test).

Because of strong species × incubation and species × stratification interaction effects (Table  3 ) species-specific analyses were also required. These demonstrated that stratification treatment significantly affected germination percentage in five out of the 20 species, either forbs or dwarf shrubs (Fig.  3 ). In E. angustifolium, Silene dioica, Vaccinium myrtillus and V. uliginosum germination percentages were higher in the ‘thick snow cover’ than the ‘thin snow cover’ treatment. Silene acaulis, on the other hand, performed better after ‘thin snow cover’ stratification (see Table  4 for mean differences in germination percentages).

Germination percentages (means ± s.e, n = 4) of the individual species in the different incubation and cold stratification treatments. See Table 2 for the incubation and stratification temperatures. Significant levels of incubation treatment (I), stratification treatment (S) and I × S interaction by two-way ANOVA are shown as follows: ns , P > 0·05; *, P < 0·05; **, P < 0·01; ***, P < 0·001. Different letters indicate significant differences between the incubation treatments at the P < 0·05 level (Tukey multiple comparison test).

Table 4.

Effects of summer warming (+2·5 °C), a reduced snow cover and the combination of both on germination (Germ) percentage and mean germination time (MGT) for each species

Warming (+2·5 °C) Reduced snow cover Warming + reduced snow cover
Species Germ (%) MGT (d) Germ (%) MGT (d) Germ (%) MGT (d)
Alnus incana n.s. −10 n.s. n.s. n.s. −10
Astragalus frigidus n.s. n.s. n.s. n.s. n.s. n.s.
Betula nana n.s. −9 n.s. +2 n.s. −7
Betula pubescens n.s. −9 n.s. n.s. n.s. −9
Betula pubescens ssp. czerepanovii n.s. −6 n.s. +3 n.s. −3
Calamagrostis lapponica n.s. −7 n.s. n.s. n.s. −7
Calamagrostis purpurea n.s. −7 n.s. n.s. n.s. −7
Deschampsia flexuosa n.s. n.s. n.s. n.s. n.s. n.s.
Dryas octopetala n.s. −8 n.s. +3 n.s. −5
Epilobium angustifolium n.s. −5 −5 +2 −5 −3
Festuca ovina n.s. −6 n.s. n.s. n.s. −6
Pinus sylvestris n.s. −6 n.s. +6 n.s. n.s.
Salix glauca n.s. n.s. n.s. n.s. n.s. n.s.
Silene acaulis n.s. −6 +19 −2 +19 −8
Silene dioica n.s. −7 −15 +10 −15 +3
Solidago virgaurea n.s. −5 n.s. +3 n.s. −2
Vaccinium myrtillus n.s. −18 −21 +5 −21 −13
Vaccinium uliginosum n.s. −9 −33 +3 −33 −6
Vaccinium vitis-idaea n.s. −14 n.s. n.s. n.s. −14
Vicia cracca n.s. n.s. n.s. +12 n.s. +12

For warming, the estimated marginal means of the ‘ + 2·5 °C’ incubation treatment were compared with the ‘control’ treatment; a negative value for MGT thus indicates faster germination in case of warming. For reduced snow cover, the marginal means of the ‘thin snow cover’ stratification treatment were compared with the ‘thick snow cover’ treatment. ‘Warming + reduced snow cover’ gives the sum of both treatments.

Significant effects of incubation treatment on germination percentage were found in S. acaulis (+15 %), V. myrtillus (+67 %), V. uliginosum (– 15 %) and V. vitis-idaea (+14 %) when the ‘optimum’ incubation treatment was compared with the ‘control’ incubation treatment (Fig.  3 ). Contrary to the other species, germination percentage decreased with increasing incubation temperature in V. uliginosum, but only in the ‘thin snow cover’ stratification treatment (significant incubation × stratification interaction; Fig.  3 ). In none of the species did a temperature increase of 2·5 °C (‘ + 2·5 °C’ versus ‘control’ incubation treatment) result in higher germination percentages (Fig.  3 and Table  4 ).

Mean germination time

The speed of germination differed significantly between the functional types (ANOVA, F5,447 = 29·0, P < 0·001). The dwarf shrubs (41 d) and the legumes (40 d) germinated significantly slower than the other functional types, and the forbs (19 d) were the faster germinating group (Fig.  1 B). Large-seeded species germinated a little slower than species with smaller seeds (linear regression, F1,451 = 4·53, P = 0·034, R 2 = 0·01).

MGT was significantly affected by the incubation and stratification treatments (Table  3 ). In general (all species combined), the germination time was shorter after ‘thick snow cover’ (27 d) than after ‘thin snow cover’ (30 d) stratification (Fig.  2 B). Further, germination was fastest in the ‘optimum’ incubation treatment (16 d), followed by the ‘ + 2·5 °C’ (31 d) and the ‘control’ (39 d) incubation treatments (Fig.  2 B). A lack of interaction between incubation and stratification (Table  3 ) resulted in the shortest MGT when ‘thick snow cover’ stratification was followed by ‘optimum’ incubation (14 d) and the longest MGT when ‘thin snow cover’ stratification was followed by ‘control’ incubation (40 d).

Analyses on individual species level revealed significant stratification effects on MGT in eleven species, belonging to all functional types except the grasses (Fig.  4 ). Whereas ‘thin snow cover’ stratification usually delayed germination, the opposite was observed in S. acaulis. The strongest effects occurred in S. dioica and V. cracca, in which germination was on average delayed by 10 d and 12 d, respectively, after ‘thin snow cover’ stratification (Table  4 ).

Mean germination times (MGT) of the individual species in the different incubation and cold stratification treatments (means ± s.e., n = 4). See Table 2 for the incubation and stratification temperatures. Significant levels of incubation treatment (I), stratification treatment (S) and I × S interaction by two-way ANOVA are shown as follows: ns , P > 0·05; *, P < 0·05; **, P < 0·01; ***, P < 0·001. Different letters indicate significant differences between the incubation treatments at the P < 0·05 level (Tukey multiple comparison test).

In all species except the legumes (Astragalus frigidus and V. cracca), incubation temperature significantly affected MGT (Fig.  4 ). Germination was slowest in the ‘control’ treatment and was faster in the ‘ + 2·5 °C’ and the ‘optimum’ incubation treatments, but in Deschampsia flexuosa and Salix glauca no difference was seen between the ‘control’ and the ‘ + 2·5 °C’ treatment. In D. octopetala, E. angustifolium and V. uliginosum, the strength of the stratification effect depended on the incubation treatment (significant incubation × stratification interaction), whereas for the other species the effects of incubation and stratification were additive (Fig.  4 ). In ten species, germination occurred at least 1 week earlier in the ‘ + 2·5 °C’ than in the ‘control’ treatment; in V. myrtillus and V. vitis-idaea MGT was even reduced by 2 weeks (Table  4 ).

Germination in a future climate

In the majority of species, the positive effect of warming (+2·5 °C) on germination time was diminished if the seeds were first stratified in the ‘thin snow cover’ treatment (Table  4 ). Eight species, though, still showed a reduction in MGT by at least 1 week. In E. angustifolium, V. myrtilllus and V. uliginosum, the advantage of faster germination was counterbalanced by reduced germination percentages after ‘thin snow cover’ stratification. Silene acaulis, on the other hand, showed both an increased germination percentage and faster germination in the scenario of warming combined with a reduced snow cover. Silene dioica and V. cracca suffered most from the future scenario: V. cracca with a strongly delayed germination (+12 d) and S. dioica with delayed germination (+3 d) combined with a reduced germination percentage (−15 %).

Germination pattern across time

To examine how germination was influenced by the different temperature steps in each of the incubation treatments, the accumulated germination percentages (averages of the two stratification treatments) were plotted over time for two species (Fig.  5 ). In B. nana, germination started during the first week of incubation and almost immediately reached its maximum percentage in the ‘optimum’ incubation treatment (Fig.  5 A). In the ‘control’ and ‘ + 2·5 °C’ treatments, germination started under June temperatures and reached maximum values under summer temperatures. Despite the strong delays in germination under colder incubation temperatures, final germination percentages were similar in all treatments. This pattern was observed in the majority of species (not shown). A different pattern was observed in V. myrtillus (Fig.  5 B) and V. vitis-idaea (not shown), in which germination occurred only after summer temperatures had been reached in the ‘control’ and ‘ + 2·5 °C’ treatments. In the ‘optimum’ treatment, germination started under June temperatures and reached a higher final germination percentage than in the colder treatments, but percentages in the ‘ + 2·5 °C’ and ‘control’ treatments were similar, after an initial delay. Figure  5 clearly shows that germination is strongly temperature regulated, with no germination until a certain temperature threshold has been reached.

Accumulated germination percentages (means ± s.e., n = 8) of (A) Betula nana and (B) Vaccinium myrtillus after different weeks of incubation in the different incubation treatments. Months indicate during which time of year the selected temperatures occur in the Abisko region.


The results demonstrate strong effects of stratification and incubation temperatures on the timing and level of germination in a large set of subarctic plant species. Although the experiment carried out did not include all abiotic and biotic changes that go along with a warmer climate in a natural environment, we believe it is justified to focus on temperature because this factor has several times been proven to be the most important environmental variable regulating the dormancy state and germination of seeds (e.g. Roberts, 1988; Vleeshouwers et al.; 1995; Probert, 2000).

In earlier studies on the influence of snow cover and freezing temperatures on subsequent germination, no significant effects were found. In a study by Graae et al. (2008), there was no difference in germination percentage between seeds of E. hermaphroditum and V. uliginosum stratified at 0·5 °C in incubators and seeds experiencing colder winter temperatures outdoors at boreal (−0·5 to −2 °C) and Arctic (mean winter temperature of −6·9 °C) sites. Similarly, Baskin et al. (2002) could not detect any difference between full- and half-snow-cover treatments on the germination of E. hermaphroditum. However, it was found that colder winter soil temperatures delayed the germination in ten species and decreased the germination percentage in four species, whereas the opposite was found in S. acaulis. The results are thus the first to demonstrate that the projected reductions in snow depth and duration in (sub)arctic and alpine regions (ACIA, 2004; Keller et al., 2005) may have important implications for seed germination, both by affecting the number of germinating seeds and the timing of germination.

Regarding summer warming, it was observed that even relatively small increases in temperature (+2·5 °C) strongly reduced the germination time in all but four species. The lack of a response in the legumes is possibly due to their physical dormancy, which is less temperature dependent than physiological dormancy (Baskin and Baskin, 1998; Probert, 2000). Contrary to expectations (Baskin and Baskin 1998; Graae et al., 2008), a positive relationship between incubation temperature and germination percentage was not commonly observed. This occurred only in three species when the optimum treatment was compared with the control treatment; otherwise final germination percentages were similar. Lower incubation temperatures thus mainly resulted in delayed germination, rather than in lower percentages. Because germination studies are usually finished after 4 weeks (for a review, see Baskin and Baskin, 1998), we think that the frequently reported lower germination percentages at lower incubation temperatures may often be an artefact of the experimental procedure. Indeed, if the present experiment had finished after 4 weeks, a similar report on the different germination percentages between the incubation treatments would have been made (see Fig.  5 A). Therefore, we stress the importance of continuing germination trials until germination percentages stop increasing.

Most species were able to germinate under June temperatures in the control treatment, suggesting that a temperature of 10·5/2 °C (during several weeks) is generally high enough for germination in subarctic species, although germination occurred much faster at slightly higher temperatures. As suggested by Körner (2003), a great functional variability in the germination behaviour of the species was found. For instance, the high-altitude species S. acaulis and D. octopetala started germinating during incubation at 6·5/3 °C, whereas V. myrtillus and V. vitis-idaea needed 12·5/4·5 °C for several weeks before the onset of germination, suggesting that species from higher elevations may be better adapted to germinate under low-temperature conditions. This contradicts the idea that seeds from higher elevations need higher germination temperatures than those from lower altitudes to avoid germination in early spring when the probability of frost is still high (Billings and Mooney, 1968; Cavieres and Arroyo, 2000). In addition, S. acaulis was the only species that benefited from the ‘thin snow cover’ stratification treatment, both with a higher germination percentage and a shorter germination time. Apparently it is especially well adapted to perform well in the harsh wind-swept habitats where it mostly grows. The protective nature of the dense cushions in which it occurs (Körner, 2003) may enable its seedlings to overcome frost and drought without much damage. The other high-altitude species, D. octopetala, did not show the same adaptation, which may explain why this species reproduces predominantly by means of clonal growth (Wookey et al., 1995).

In the majority of species, the combined effect of a reduced snow cover and summer warming resulted in faster germination, but four species were not affected and two species showed slower germination. The effect on germination percentage was positive in one species, negative in four species and not significant in all others. In subarctic ecosystems, the short growing season (approx. 3 months in the tundra near Abisko; Molau et al., 2005) is a major barrier for the survival of seedlings because it constrains the period during which seedlings need to attain a critical biomass and acquire resistance to freezing to withstand the harsh and long-lasting winter conditions (Maruta, 1994; Stocklin and Baumler, 1996; Schütz, 2002). Seedling mortalities in cold climates are usually high and 12-month losses often exceed 50 % in large-seeded and 99 % in small-seeded species (Jolls and Bock, 1983; Körner, 2003). This implies that even relatively small reductions in germination time may have substantial effects for recruitment from seed by improving the chance of seedlings to survive the following winter (Chambers, 1995). Consequently, the present results suggest that a warmer subarctic climate is likely to be beneficial for seedling establishment in most species, even though colder soil temperatures during winter may diminish the positive effect.

The species that may benefit most are trees (A. incana and B. pubescens), (dwarf) shrubs (V. myrtillus, V. vitis-idaea and B. nana) and grasses (C. lapponica and C. purpurea), in addition to the forb S. acaulis. These species showed considerably earlier germination (>1 week), sometimes combined with an increased germination percentage (especially in S. acaulis) when stratified and allowed to germinate under future conditions. Vaccinium myrtillus, however, may only improve its recruitment in more protected, snow-rich places (strong negative effect of colder winter temperatures), but its germination percentage is expected to increase substantially if summer temperatures increase >2·5 °C. It has already been documented that dwarf shrubs, grasses and trees will expand in Arctic and subarctic areas (Sturm et al., 2001; Kullman, 2002; Dullinger et al., 2004; Tape et al., 2006) and that they are predicted to perform better in terms of productivity in a warmer climate (Parsons et al., 1995; Arft et al., 1999; van Wijk et al., 2004; Walker et al., 2006). Thus, the functional types that are expected to benefit most from warming in the adult stage may also benefit most in the recruitment phase, possibly resulting in a positive feedback.

The species in which germination was most negatively affected by a warmer climate were three forbs that currently occur in subarctic birch forest (E. angustifolium, S. dioica and V. cracca) and the dwarf shrub V. uliginosum. Seeds of these species apparently suffer greatly from below zero temperatures during winter, resulting in less and slower germination. However, they performed well after stratification under mild winter temperatures, so, similarly to V. myrtillus, their expansion is expected to be restricted to snow-rich places, for instance close to shrubs which trap and hold snow during winter (Sturm et al., 2005). We can, however, not explain why the combination of ‘thin snow cover’ stratification and ‘optimum’ incubation was extremely unfavourable in V. uliginosum.

The observed species-specific responses may have consequences for the patterns of species migration due to warming. In some species, seedling recruitment will improve more than in others in a warmer climate; therefore not all species will be able to shift northward or upslope at the same pace. This may result in plant communities without previous analogues (Davis, 1989; Kullman, 2002) or in shifts in dominance within existing communities (Kelly and Goulden, 2008). In any case, the importance of species-specific reactions, also during the recruitment stage, should be taken into account in models that forecast population dynamics and future species migrations.

Seed germination does not appear to limit northward range expansion in the three boreal tree species of the present experiment. Alnus incana, B. pubescens and P. sylvestris all germinated equally well (similar germination percentages) in the colder stratification and incubation treatments as in the warmer ones. Moreover, they did not germinate significantly slower under current subarctic conditions than the dominant tree species in the Abisko region, B. pubescens ssp. czerapanovii. The current distribution of these species is thus most likely limited by other processes than seedling recruitment.

To conclude, it was found that colder winter soil temperatures are likely to result in lower germination percentages and slower germination in about half of the subarctic species. Warmer summers, on the other hand, reduce the germination time and may sometimes increase the number of germinating seeds. The combined effect of both is beneficial in most species and therefore seedling establishment was expected to improve in a future warmer subarctic climate.

Planting for Pollinators: Establishing a Wildflower Meadow from Seed [fact sheet]

This fact sheet received the American Society of Horticultural Science 2019 Award for Outstanding Fact Sheet – Extension Division.

Native bees and other pollinators are essential to the successful production of many fruit and vegetable crops and the reproduction of many plant species in our surrounding environment. Wildflower meadows and gardens are extremely valuable habitat, providing floral resources, nesting sites and a protected environment for hundreds of bee species, moths and butterflies, and other insects. Many birds, bats, small mammals and some amphibians also thrive on the food and shelter that a meadow ecosystem provides.

Meadows provide many important ecosystem services including infiltration and filtration of stormwater, carbon storage, nutrient recycling, soil building, and provisioning of food and shelter for biodiverse communities of flora and fauna. By establishing native perennials and grasses in a dense and diverse meadow planting, property owners can enjoy the beauty of a succession of flowers and plant forms and experience a renewed connection with nature. Done properly, wildflower meadows are ecologically-friendly landscape components that, once established, have minimal maintenance requirements.

Soil testing can provide you with useful information regarding pH and organic matter content, but wildflowers generally prefer low fertility sites. Determine whether the soil tends to be wet or dry, and if the site gets full sun, filtered sun or shade.

Choosing a Site

Not all wildflowers are suitable for all conditions. A site with full sun and good drainage is ideal for many species, but partial shade and/or wet areas can be tolerated by many others. Consider your site and soil conditions carefully in order to select an appropriate wildflower mix.

It’s best to start in a small area, but consider 400 square feet to be a minimal size for a wildflower meadow – this space can support a good diversity of wildflower species. Some types of wildflowers get quite tall and may tend to lean or flop, but they will help hold each other up if planted densely in an area where they will not interfere with walkways or other landscape features. A wildflower meadow is informal by nature, and can be a bit wild and untidy looking at certain times of the year, so locate it where it will be viewed from a distance of several meters or more. For a neater, more designed meadow look, purchase small transplants instead of starting from seed, and plant in intentional groupings as in a garden.

A place where bees can come and go safely with little disturbance or exposure to pesticides or other household chemicals is ideal. Many native bees need patches of bare soil nearby in which to make their nests; others will nest in small holes in dead wood or stems, in cavities in stone walls or in leaf litter or debris piles. These features are often found along the edges of fields or woodlands and should be preserved. Some people build or purchase bee boxes or bee houses to encourage mason bees and other solitary bees to nest near their crops or gardens.

Seed Mixes and Species Selection

There are many wildflower mixes available from reputable seed companies1 , or you can design your own mix. Pre-made mixes may be convenient, but must be selected carefully to avoid paying for species that are unlikely to be successful in New England, or that might be overly aggressive. Less expensive mixes frequently contain a higher proportion of grasses than desired for good pollinator habitat.

Knowing your site characteristics (wet, medium or dry soil and full sun, filtered sun, or shade, at a minimum) is essential to understanding which species will thrive on your site and create a mixed meadow that knits together in a mosaic of colors and textures.

An ideal meadow mix will provide a continuous sequence of bloom from a dozen or more native perennial wildflowers (a.k.a. forbs). A few native warm-season grasses should be included to create habitat and shelter for many organisms, and to create dense cover that suppresses weed growth. Some mixes contain annuals, which will give you firstseason color but are generally not native species and not sustainable over time. You are better off in the long term to spend your money on perennial species. By including black-eyed Susan, and perhaps dotted horsemint, you can still get some blooms the first season to satisfy the need for color and provide some flowers for bees.

A suggested wildflower mix for medium to dry sites in full to partial sun is given in Table 1. This list is composed of reliable species that have performed well in research trials at the NH Agricultural Experiment Station in Durham and elsewhere around the state. Not every species will thrive (or even survive) on every site, but most are widely adaptable.

Seed mixes can be customized to meet your objectives and budget. The cost for the seed mix in Table 1 will run an average of $60-80 per 1000 square feet, if seeded at a rate of 20 lbs per acre (0.5 lbs/1000 square feet). If budget is not a limiting factor, you might want to add seeds of native lupine and butterfly weed, both of which require very dry soils. If your site tends to be wet, a more appropriate mix would contain cardinal flower, blue vervain, Culver’s root, golden Alexanders, boneset, swamp milkweed and Joe-Pye weed.

You can either purchase seed packets and mix the seed yourself, or a wildflower seed company may mix it for you on request. Order seed early in the summer to avoid shortages of popular mixes or species. Ask to have the (large and fluffy) native grass seed packaged separately from the (mostly tiny) wildflower seed. Store the wildflower seed in the refrigerator until planting time. The grass seed would ideally be refrigerated as well, but if there isn’t room for it, keep it in another cool dry place.

UNH custom mix of reliable species, suitable for sunny sites with medium to dry soils and a pH of 5.5 or above. Suggested seeding rate is 0.5 lbs per thousand square feet of area.

Wildflower Species, Common Names, Percent of mix by weight
  • Aguilegia canadensis, Red Columbine, 3%
  • Asclepias syriaca, Common Milkweed, 3%
  • Chamaecrista fasciculata, Partridge Pea, 8%
  • Coreopsis lanceolata, Lanceleaf Coreopsis, 3%
  • Echinacea purpurea, Purple Coneflower, 7%
  • Echinacea pallida, Pale Purple Coneflower, 11%
  • Eutrochium purpureum,Purple Joe-Pye Weed, 1.5%
  • Heliopsis helianthoides, Oxeye Sunflower, 2%
  • Monarda fistulosa, Wild Bergamot, 0.50%
  • Monarda punctata, Dotted Horsemint, 0.50%
  • Oligoneuron rigidum, Stiff Goldenrod, 0.50%
  • Penstemon digitalis, Foxglove Beardtongue, 1.5%
  • Ratibida pinnata, Yellow Coneflower, 3.5%
  • Rudbeckia hirta, Black Eyed Susan, 2%
  • Solidago speciosa, Showy Goldenrod, 1%
  • Symphyotrichum novae-angliae, New England Aster, 1%
  • Symphyotrichum laeve, Smooth Blue Aster, 2%
Grass Species, Common Names, Percent of mix by weight
  • Elymus canadensis, Canada Wild Rye, 10%
  • Shizachyrium scoparium, Little Bluestem, 30%
  • Sorghastrum nutans, Indian Grass, 10%

Site Preparation

Successfully establishing a meadow from seed is a three-year process, with the first year devoted to good site preparation. This isn’t the fun part but eliminating competitive weeds before you plant is essential to long-term success. How to get started depends on the beginning site conditions and what materials and methods you decide to use. The following methods have been developed based on research and demonstration plantings in New Hampshire and are appropriate for the Northeast and other areas with similar climate patterns.

Smothering with black plastic excludes light from the underlying weeds, preventing photosynthesis which is essential for plants to survive over time. Although black plastic absorbs some heat during the day, soil temperatures underneath do not get high enough for long enough to kill weed seeds. Solarization, in contrast, acts by trapping solar radiation and converting it to heat underneath clear plastic sheeting. Extreme temperatures for long durations can reduce viability of sensitive weed seeds, although the results are inconsistent in our climate.

What are You Starting with?

Rough turf or lawn areas – It is essential to completely kill existing grasses and other perennial weeds in a turf area before planting wildflowers. A full season of site preparation is critical to success, because young wildflower seedlings stay small and low to the ground their first year of growth and are not able to compete against more vigorous weeds. Perennial grasses, such as our cool season lawn and pasture grasses, spread from strong underground roots and rhizomes and are especially troublesome when trying to establish wildflowers. These and other perennial weeds can be effectively killed by a process called “smothering” during the course of the summer prior to planting wildflower seeds. Steps to take:

  1. Mow the area as short as possible once or twice after it greens up in the spring. Scalp it! Then, rake off any excessive organic matter to create a smooth surface. Leaving a light layer of clippings is okay. Do not till the soil.
  2. Lay sheets of thick (4- or 6-mil) black plastic over the entire area, overlapping the edges by about a foot if you use more than one sheet or roll of plastic. Bury all the outside edges with soil, and/or hold the plastic down with rocks, cinderblocks, bricks or other available materials. The objective is to exclude light from the grass and weeds trying to grow under the plastic. Weeds without light are unable to photosynthesize and will eventually run out of energy and die. Any seeds that germinate under the plastic are likewise unable to survive for long.

Smothering is an easy, inexpensive, no-till method for preparing small to medium size areas. Alternatively, in turf areas you could remove the top layer of sod with a sod cutter and then apply one of the site preparation methods described for cultivated soil, below

Use herbicides with caution. Follow all label instructions including the use of protective clothing. Do not spray when weeds are in bloom or bees are present.

Cultivated soil (such as an agricultural field or garden) – A piece of land that has been recently cultivated for crops may appear to be relatively weed-free; however, there is usually a reserve bank of weed seeds lying dormant in the soil. Eliminating seedlings as they appear (and before they set seed) will diminish the weed seed bank over time.

Four options for reducing the weed seed bank are presented below. Any of these strategies should be implemented for the entire growing season prior to planting the meadow mix. The first two methods tend to be most effective for annuals and some perennials reproducing from seeds, but are less effective where perennial grasses are already well-established; if that is the case, use of herbicides or the smothering method described above is more effective.

  1. Repeated tillage or cultivation. If weed pressure is low, manually pulling or hoeing to kill germinating seedlings may be sufficient, but for large areas and/or heavy weed pressure, a mechanical cultivator is recommended. If perennial grasses or deep rooted weeds are present, the area should be tilled deeply first, then shallower cultivations used subsequently. Cultivating at a shallow depth should suppress germinating annuals and some broadleaf perennials but is less effective on clover and grasses, especially those coming back from root pieces or rhizomes. Repeat the operation whenever a flush of weeds appears.
  2. Cover crops. Planting a dense summer cover crop will suppress weeds by shading and competing for space. Buckwheat is a good choice and will bloom and provide floral resources for bees during the summer. Mow or roll it at the end of the bloom period to prevent seed set; any live tissue remaining will be killed by freezing temperatures in fall. A late season crop of oats is also a good option, as oats will be killed by winter temperatures. Cover crop residue must decompose before seeding wildflowers, however, so fall planting is not feasible. Rake off debris and smooth the soil surface before seeding the following spring, but tilling is not recommended as it will bring up more weed seeds.
  3. Herbicides. Two or three applications (early season, mid-season and possibly early fall) of a nonselective systemic herbicide is a highly effective method for killing actively-growing annual and perennial weeds. There are alternative “burn down” and/or organic products which may be used, but these are not translocated in the plant to kill roots and rhizomes; therefore, more applications are needed to eliminate weeds as regrowth occurs and/or new seeds germinate throughout the growing season. Do not till the soil after using herbicides in order to avoid bringing up more weed seeds.
  4. Solarization. This method involves covering bare, moist soil with clear (not white) plastic from mid-June through mid-September or longer. Effective solarization depends on trapping solar radiation as heat, raising the soil temperature high enough to kill weed seeds. Solarization is most reliable in hot, arid climates, but may yield some success if the site is fully exposed to sun and the edges of the plastic are buried to effectively trap heat. Use 4- or 6-mil UV resistant plastic, such as growers use to cover high tunnels or greenhouses. Used plastic is okay but will develop tears and holes more quickly from photo degradation or from animals such as deer walking across the surface. Use heavy-duty clear repair tape to seal up any tears and holes which occur as soon as possible, or the openings will act as vents and make the solarization process less effective. There are conflicting reports on the effectiveness of soil solarization on weed suppression in northern climates and results may vary from year to year. In a cloudy, cool season it is apt to be unsatisfactory.

Forest/woodland sites – recently logged or cleared woodlands usually have low soil pH and nutrient levels, but still can become successful meadow areas. Stumping, grading and/or excavating and raking is usually necessary. The period of time between clearing and seeding may be bridged with an annual grass crop or cover crops. Watch to see what weeds (or tree sprouts!) emerge during this time and use herbicides and/ or mechanical controls as needed. Cover crops which are winter killed, such as oats or buckwheat, will help smother new weeds that come up during the growing season, but essentially prevent fall planting, as they must first decompose or be removed. It is best not to till, in order to prevent new weed seeds from being brought to the surface, as well as rocks and roots.

An effort should be made to select species that are tolerant of acid soils. However, if pH is very low (<5.5), it may be prudent to make an application of lime and other amendments as recommended by a soil test. Incorporation of amendments by tilling creates disturbance and may result in high subsequent weed pressure, but may be needed to improve growth on poor, acid sites. Tilling may bring up roots, rocks and other items, requiring more raking and smoothing before seeding.

Other disturbed sites – construction sites or other sites which have been excavated, graded and/or filled with soil from off-site present a special challenge because conditions are hard to predict. Gathering as much information as possible will help you select appropriate meadow species for planting. Have the soil tested for soil texture, pH, nutrient levels, and organic matter. Once the site work is done, observe water movement and do a percolation test to check drainage. Using all this information, select a mix of species most suitable for the conditions.

To determine what the potential weed pressure is, observe what types of weeds come up over time. To expedite this process, you can fill several nursery pots (or other large containers with drainage holes) with soil and water them regularly to see what weeds emerge. Based on your observations, determine whether there is a significant weed bank, in which case you should proceed as if it was a previously vegetated area, or if weed competition is minimal and you can plant soon.

Planting – When and How

After a season of site preparation, you are ready to plant! Remove plastic or other mulch materials and rake off any loose tree roots, leaves, cover crop residues, etc. to prepare a clean seed bed. Fall is a good time to seed, as wildflower germination will be enhanced by exposure to cold temperatures and damp soil during the winter. The fall planting season in northern New England extends from late September through early December, depending on the year. A safe strategy is to aim for mid- to late-October, whereas November weather is unpredictable and snow could cover the ground at any time. If that happens, or if there are other reasons you choose to plant in the spring, store your seed in the refrigerator or other cold (35-40°F), dry place for the winter and then plant it as early in the spring as possible.

Broadcasting (spreading seed by hand) is the preferred method for small areas. A carrier such as vermiculite or sand is needed to “bulk up” the volume of material to be distributed

You might want to practice distributing the moistened carrier without any seed at first, to practice getting the right amount evenly distributed over the desired area. Use a broad sweeping throw for each handful, much like feeding the chickens.

When it’s time to plant, measure the area to be planted and perhaps divide it into smaller subplots of 400-500 square feet each to make seed distribution more precise. Calculate the amount of each type of seed (grasses and wildflowers) you need for each subplot and set it aside.

Mix the seed with vermiculite or other carrier for one subplot at a time in a plastic paint bucket or similar container. Start with the dry carrier then add small amounts of water at a time, stirring until it is slightly damp but not wet. Now you can mix in your seeds; the small seeds will stick to the carrier particles, keeping everything well-mixed for distribution. Add the wildflower seeds first, then the grass seeds, adding more water or vermiculite if needed. The precise amount isn’t important, but using 0.5 to 1 gallon of the carrier per 500 square feet area is adequate. Wearing latex gloves will keep the seed from sticking to your hands.

For each subplot, apply half the amount of seed while walking back and forth in one direction, then repeat in the other direction, as shown in the diagram below. Using a light-colored carrier such as vermiculite allows you to see how evenly you have distributed the mix on the soil surface. Scatter any remaining seed where needed.

Raking lightly with a metal lawn or leaf rake after broadcasting helps work the seeds into the soil, but rake only ¼” deep so you do not bury the tiny wildflower seeds. If the soil is firm and level, skip raking and go straight to rolling the area with a lawn roller, or a cultipacker, which will press the seed into the soil. Good seed to soil contact is essential for holding the seed in place over the winter and helps keep the seedlings from drying out once they germinate in the spring. And finally, a thin layer of clean straw (one bale per 1000 square feet) distributed lightly over the top helps keep the seed in place.

If seeding a large area (of several thousand square feet), you might want to try a mechanical seeder. A slitseeder or seed drill can be used on smooth level sites, such as an agricultural field, but most people don’t have access to that equipment, which may also be difficult to calibrate. A whirlybird-type broadcast lawn seeder or chest-carried crank seeder may work satisfactorily, but it is hard to keep the seed well-mixed and feeding properly; the tiny wildflower seeds tend to fall to the bottom and get used up first. Calibrate your seeder with a dry carrier (such as sand or rice hulls) before mixing in any seed. Then mix small batches of seed with the carrier, use half the amount going back and forth in one direction, then repeat in a perpendicular direction. In this case, do not add water to moisten the carrier, as it will prevent it from feeding through properly.

What to Expect

Year 1 – is the season for site preparation, an essential but not very attractive process. Time and effort spent this year will provide a clean seedbed to be planted and mulched in the fall or following spring. Skipping the site preparation process is sure to result in failure over the long run.

Year 2 – you will most likely be disappointed in how your meadow area looks the first season after planting. Patience is the key this year. You should see wildflower seedlings germinate and emerge as the soil warms up in the spring, but it’s hard to tell the wildflowers from the weeds at this point. Some wildflowers won’t even germinate for 2-3 years following planting, and most grow low to the ground the first season. If you haven’t done a great job of preparing the site and killing existing vegetation, weeds will grow up quickly and can easily smother or shade out the wildflowers. Hand-weeding may disturb germinating wildflower seedlings, so is not recommended. Consider mowing in mid-summer at a 4-6” mowing height, whacking the weeds back but going right over the top of most wildflower seedlings. Few wildflowers will bloom this year anyway, as they are devoting their energy to growing strong roots and shoots rather than flowers and seeds. Black-eyed Susan is the exception to the rule, so be sure to include it in your seed mix to provide cheerful yellow flowers this year. It will even recover and re-bloom after that mid-summer mowing.

Crabgrass is a special challenge on some sites. A thick blanket of crabgrass can smother out germinating wildflowers, and is not sufficiently managed by mowing. There are few options for controlling crabgrass other than use of a post-emergent selective grass herbicides that can be sprayed over-the-top of wildflowers. One application to actively growing crabgrass before it goes to seed will effectively kill it, reopening the area to allow light to reach the underlying wildflowers. As with all herbicides and other pesticides, follow label directions carefully, and consider whether hiring a licensed applicator is required or prudent for the situation at hand.

Year 3 – If they survived last year, the wildflowers will emerge quickly in the spring and grow much faster and larger this year. Most weeds are slower to get started and pose much less of a problem this year, often being out-shaded and out-competed by a dense wildflower mix. By late June, you should see some flowers on coreopsis and columbine, if they are in your mix, followed by foxglove beardtongue and blackeyed Susan. In mid-summer, wild bergamot (bee balm) and oxeye sunflowers will bloom prolifically, and perhaps a few yellow and purple coneflowers will appear. Wild Rye shoots up and provides a pleasant contrast with its distinctive seed heads. Other warm-season grasses may still be slow and inconspicuous.

Year 4 and beyond – You and the bees will reap the rewards of your efforts, enjoying a dense, diverse mix of colorful wildflowers from spring through late fall, when goldenrods and asters provide a fall feast for bees. Black-eyed Susan, coreopsis and a few others will diminish in numbers and tend to migrate to the edges of your meadow. The mid-summer meadow buzzes with bees and other insect pollinators, and birds reap the benefit of bugs and seeds to eat. Milkweed finds its place in the meadow, and monarchs feast on the nectar of many meadow flowers before laying their eggs on the undersides of milkweed leaves. Warm-season grasses fill in areas where the wildflowers are less dense, providing clumps that shelter ground-nesting bees and other creatures.

These photos show the growth and development of the same meadow planting over time.

Long Term Changes and Maintenance

Once you have an established meadow, there is little you need to do. Monitor for tall and aggressive weeds such as sumac, pokeberry, purple loosestrife and bittersweet vine, removing them by hand in the fall when you can pull or wrench the roots out. Try not to disturb the soil any more than necessary in the process, because disturbance creates openings for future weeds and invasives. You can continue to edit the meadow by adding plants to sparse areas and preventing some of our volunteer wildflowers (a.k.a. weeds) from taking over. Some to watch for include daisy fleabane, evening primrose and common mullein. These are all good pollinator plants but may outcompete some of the more desirable species you planted, reducing diversity. Cutting them back before they set seed will help keep them in check.

Once the meadow is finished flowering and freezes kill the last of the asters to the ground, consider the beauty in the structure and golden colors of standing seed heads and grasses, which the birds will appreciate well into the winter. If you need to tidy up, mow the meadow down in November or leave it stand until early spring. Mow high (6-8” or higher) and wildlife will continue to nest and forage in the meadow through the winter and spring. Mowing every year is not required; its primary purpose is to discourage woody shrubs and trees from taking over. If you have a large meadow area, consider mowing only one-third or one-quarter of it each year, leaving the rest for winter habitat.

The meadow is an ever-changing landscape. Weather variations, soil conditions, and wildlife will determine which wildflowers and grasses become dominant and which fade away over time. In a wet year, plants like Joe Pye-weed and cardinal flower may flourish, while in a dry year bergamot and coneflowers may abound. Take a step back and enjoy the changes – a meadow is a process, not a product!

Acknowledgements: This work was funded by the USDA National Institute of Food and Agriculture Hatch Multistate Project 1010449, with additional support from the Anna and Raymond Tuttle Environmental Horticulture Fund and the NH Horticultural Endowment. The author gratefully thanks Amy Papineau, Sean Fogarty, Sam MacNeil and the farm crew at the AES Woodman Horticulture Farm for all their help.

See also  orange valley og seeds