Weed Ecology - Science topic

Hello Susan; I'm very curious to know what finally came of your question. Did you learn what you needed to know? Were you able to act on it? Best regards, Jim Des Lauriers

Cheatgrass (Bromus tectorum) is an invasive plant. Please see the below given RG links.

Please take a look at the following RG link.

This post raises several interesting questions:

1) I wonder how much allelopathy contributed to the competitive exclusion (or other recruitment failure) of many planted species. In any case, it is indeed common for many seeded species to not successfully establish.

2) I think we are setting too high a standard for "success", when we say that every seeded species that does not establish is a restoration "failure". I think it is common to add species to a seed mix to maximize the chances that some good diversity is established, not that every species will. But again, I agree that restorationists are often disappointed with how few species establish, and consider this a problem.

3) Aggregated seeding (including "strip seeding") is indeed being explored as means to increase restoration success of subordinate plant species. See:

Porensky LM, Vaughn KJ, Young TP (2012) Can initial intraspecific spatial

aggregation increase multi-year diversity by creating temporal priority?

Ecological Applications 22:927–936

Yurkonis KA, McKenna TP (2014) Aggregating species at seeding may increase

initial diversity during grassland reconstruction. Ecological Restoration

32:275–281.

Although these methods have shown some short-term benefits, their long-term benefits are not clear. See:

Young DN, Porensky LM, Wolf K, Fick S, Young TP (2015) Burning reveals

cryptic plant diversity and promotes coexistence in a California prairie

restoration experiment. Ecosphere 6:81

Young, T.P., K.L. Stuble, K.L., J.A. Balachowski, and C.M. Werner. 2017. Using priority effects to manipulate competitive relationships in restoration. Restoration Ecology 25:S114–S123.

Dear Ankur,

Please have a look at these useful links.

Good luck!

The measuring of the shift in weed populations is done be taken 4 to 8 sample grids which are tossed randomly into the field for testing. Harvest all the weeds and crop in the grid. Identify and count the species and then weigh the fresh weight. Put the separated material into an oven to dry and take a dry weight after 48 hours when dry weight is constant. The change in number and weight of weeds from one year to another in the same field and area gives a measurement of the shifting in the populations. Good luck in your project. 

thank u @Andrew Paul McKenzie Pegman 。I were doing work on field last  5 months,so i had no chance to reply u。I am puzzling about its seed，could you show me some seeds ？thank u very much

It is interesting to know that the sustainability of cropping systems can be increased by introducing a cover crop, provided that the cover crop does not reduce the cash crop yield through competition. The cover crop may be sown at the same time as a cash crop in the crop rotation.

Please check the PDF attachments.

I'm not an ecologist but what happened in your control that, e.g. had the enzymes and either no stubble or an artifical, non-degradable 'stubble'?

Hi

There is a registered microbial herbicide against Cuscuta spp. (dodders), which includes Colletotrichum gloeosporioides f. sp. cuscutae. It registered in China under the name of Luboa S22.

As the plant is evergreen or may only lose some leaves in unfavorable conditions, it is clearly a phanerophyte. However, you might want to keep in mind that Raunkiaer's life forms are mainly created for (highly) seasonal (esp. temperate) floras. Tropical taxa often don't fit or differ extremely in form (cacti, trees, evergreen vines, epiphytes are all phanerophytes!) although being in the same Raunkiaer class. That's why most species are not put in one of those categories. There are other life form classifications though.

It has several ways to test it. You can search on isolation and identification of plant extract.

There a lots of different ways to randomly distribute traps (also depending on trap size) for this purpose, which mostly work well. One important point to look at is which ecosystem you are working in. This should help to define trap size.

The plot size does not necessarily have to be the same. However, the number of traps you set in each plot should be equivalent to the surface area, i.e. you should have the same number of traps/surface area in all plots.

Dear Sheik.

If you are interested in the seed rain per tree you would not need to take trap density into account. You would just use a trap under each tree and refer to the seed-rain per tree. In that case you should include tree crown size as a covariate.

However, I would say, that still the most interesting question is related to the seed rain per area unit due to the following facts:

* When working with seed dispersal the usual interest is in the number of seeds dispersed per each seed producing tree (not to be estimtated by traps).

* Appart from that the number of seeds deposited on an area seems to be ecologically more relevant than the seeds deposited under individual trees. But again, that may not hold in your case, depending on the background and context of your research.

Anyway, I would say what you present as an objective is not the ultimate objective, but rather what you intend to do. The most important question (fundamental when designing your study) is what you want to do that for...the underlying question.

In general, and without that detail, and given the weeded area is unknown, I would chose one plot at each of the habitat types. You have to be specially carefull to ensure that both plots have the same size and are in the core area of both patches (to avoid edge effects, which would be greater in the un-weeded area). You would then install an equal number of traps at both plots, each one under a randomly selected tree.

The most direct and correct response variable would be number of seed per trap. Given the design you don't need to calculate the number per ha to compare both plots. 

With that simple desing you could use a simple GLM to compare both plots, where each trap is a replicate and plot (type of patch; weeded vs un-weeded) is a fixed factor. It would be similar to a t-test, but using a Poison distribution instead of a normal distribution (with a log link-function). An alternative is to use a Kolmogorov-Smirnov or a Mann-Whitney U non-parametric test, but probably GLM is more powerful.

These statistical test will tell you if there is any difference between both plots. If there is any difference you would calculate a mean density of seeds per ha for each plot, otherwise you would pool both plots to calculate a unique density of seeds.

However, beware of the lack of replicates for the treatment. Your work will only tell you about the difference among those particular plots...not about weeded and un-weeded patches in general. You would need additional weeded and un-weeded areas to obtain such a general result.

Best regards,

       Asier

Hi,

In the Caribbean this plant species is considered sometimes as an alien species, sometimes as a native species (!), but it seems that nobody consider it a problem !.  A great number of habitats are now invaded with this plant. Often, people introduce this plant on small ponds, gardens and other standing water habitats.  People consider it as a beautiful plant that increase attractivity of ponds. We do not know any factors really helpful to limit its growth except manual withdrawal. I don't know of any tests carried out in this region against this species.

Since this species is not a problem for boats or for other commercial activity in the West Indies there is no policy considering E. crassipes

Ponds colonized by this species are also the only habitat, in the West Indies, for the dragonfly Miathyria marcella.

Best

Mr Shergill and Mr Jalaludin, I will collect some seeds of Elleusine indica from some locations. Would you like give me advice? stage seed or age of eleusine that can be collected ? thank you

looks to me like birds nest fern.

Hi Kathryn, I think you are right in general: birds of any sort are reluctant to abandon a good food source, as long as they have places available for roosting, nesting, etc.  Bell Miners are usually found in shrubby forest, and they use shrub thickets for roosting and nesting (though I know they often nest in eucalypts too, and suspect they may roost there sometimes as well).  I've suggested shrub removal as a possible way of deterring Bell Miners, and have seen places where shrubs have been removed (by fire and by grazing) and Bell Miners have abandoned those areas.  I would not expect that to happen on every occasion.  But I do believe that Bell Miner colonies need to be mobile (or the psyllids they sequester may end up killing the host trees) and removal of understorey may sometimes provide a useful trigger for them to move.  A lot depends on scale: if there are still good shrub thickets nearby for roosting and nesting, the Bell Miners may continue to use the area from which lantana has been removed.  If not, I think you might find they do move.  I'd be very interested to hear what you find!  My email is richard.loyn@bigpond.com, and I'd be happy to communicate further on this.  Cheers, Richard.

I compared the grass with Brachiaria distachya, they looked much alike. But no record of B. distachya existed for Nigeria. I have checked through literature, even those by Hutchinson & Dalziel (Flora of West Tropical Africa) and Nigerian Grasses by Joyce Lowe, but did not find any record for this species. 

After reflexion, it's a Podophyllaceae, like Bongardia chrysogonum ; see http://www.avonbulbs.co.uk/s/bongardia-chrysogonum/52/Product.aspx

It looks like corn spurry to  me. (Spergula arvensis L.). It is a fairly common weed in the United States and has been recently identified in Alaska.

The most important thing would be to test the crop of interest to see if it is susceptible to injury.

Hi Eajaz,

We collect seed for susceptible checks from non-agricultural areas with no history of herbicide use (e.g. public park, home garden, roadsides, other natural areas, etc.). This seems to work well. I think there are a variety of approaches you can use here, as long as you can justify that the check should be susceptible, I think you should be good.

Scott

You seem to have a very good understand many of the limitations with visual phytotoxicity assessments. Additional problems include the issue of many researchers not doing this assessments blind (they take a plot plan with them and know what plots they are rating) so the potential for bias is VERY high. My question would be why use them at all? Because of these problems most journals will not allow publication of papers without additional truly quantitative data such as biomass and yield. Because of this we will use them sometimes as a semi-quantitative way of describing herbicide phytotoxicity but I am not sure of there utility for anything beyond that. However for those that are screening massive amounts of herbicide plots they may be a necessary tool. Looking at other disciplines that have done a better job may be informative - plant pathology in particular has been much more rigorous in there development of scales for plant pathogen symptoms.  This approach probably could be transferred to contact herbicides fairly well. The issue always will be that there are large differences between herbicide MOA and plant species. In addition for many MOA most obvious symptomatology is the cessation of plant growth that requires the comparison of the untreated plots with treated and therefore removes part of the blind rating. So is this impossible - no, but I think one has to consider if the data will really be that much better. 

Yordanys, there are a couple of good checklist on weeds for a couple of Neotropical countries. You can get Villaseñor & Espinoza-García 2004 in Diversity & Distribution 10:113-123 for Mexican weeds. Since many weeds are widely distributed you can also try this site http://www.iewf.org/weedid/iewf_front_id.html for Australian plants.

Weed seed bank study can be carried out by putting an enclosure in a particular site, collecting soil samples from this enclosure, putting this sample in a tray and allowing seeds to grow and keeping a count of the weeds emerging. Record can be kept on different weed species and time of emergence etc.

Yess..... yes you can be quantify the losses.... that is in terms of loss in aesthetic value of a property (Compare the price of a weed infested property in your area, it can reduce price by up to 15%).... Weeds can also interfere or compete with flowers, shrubs, turf and other plants in your landscape... this can lead to death of the desired plants and I am sure you can put a price to that....

The cost of controlling weeds in your landscape is another quantifiable parameter...

Again, a landscape that is infested with weeds losses aesthetic value, becomes unsightly, and if it is for commercial purposes you will end up loosing clients, translating to  opportunity costs  and loss of revenue....

So depending on your circumstances and objectives, you can quantify the losses caused by weeds in your landscape.....

The term "weed" can be applied to almost any plant, depending on the situation and the perspective of the person referring to a particular plant as being a "weed".  For example, glyphosate-resistant Zea mays could be a weed if it is found growing and competing with glyphosate-resistant Glycine max. 

Crop-weed associations could be due a number of factors: similarities in growth habit, life cycle, management practices, etc. 

Most weeds tend to be herbaceous because this type of plant tends to have the traits of a fast-growing colonizer that can utilize available resources after a disturbance.  Woody species can be "weedy", but this type tends to appear in later stages of succession. 

Sampling weed species largely depends on several factors; including cropping pattern, size of plot or area covered, etc. The most common technique is using ''quadrant'. This can be temporary or permanent (fixed) or movable quadrant. Dimension may vary depending on some of the factors mentioned above. I usually use 0.25m^2' or 1m² for sampling weeds in experimental plots. All weed species within the quadrant are collected, counted and identified according to species type.  I also usually make two or three throws of quadrant per plot. Total count per specie type and for all three throws are added together to give relative abundance of each specie per treatment. In addition, I also usually determine % weed  cover, ease of control and level of aggression per weed specie type. I hope this information can be helpful.

SMKanteh,

Njala University

Dear Jacob，thank you for your idea. LIU

climate of one year may be not enough, as climate is changeable, it is better to take in consideration the climate of several years. Of course, there is a relation between distribution, growth performance, etc of your studied weeds.  

also

Weed control programs in previous years?

Which herbicides did apply previous years?

Which species are resistant to herbicides?

this could be a good start

Sorry the title is in english: "purple jewelweed impatiens glandulifera royle in the ojcow national park"

The pregerminative conditions that I´ve worked with are: 30 min in 96% H2SO4 and wash with water for 5 min. Germination conditions are: 20 ºC/ 30 ºC, 12/12 h in complete darkness.

Hope I've been helpful!