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# Morphometric Analysis - Science topic

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Questions related to Morphometric Analysis

Submit your abstract for the Symposium organised by Ivana Rešetnik and me at the XX International Botanical Congress in Madrid (21-27 July 2024)!

The deadline for abstract submission is

**November, 30 for oral presentations**and February 1, 2024, for posters.Here you can find the submission guidelines: https://ibcmadrid2024.com/index.php?seccion=scientificArea&subSeccion=abstractGuidelines

Link for abstract submission: https://ibcmadrid2024.com/index.php?seccion=scientificArea&subSeccion=abstractSubmission1

The general link to the congress: https://ibcmadrid2024.com/

Our symposium is the n.

**13**Hello, I am a dentist doing my research to graduate. I am measuring a bone structure of the skull in adolescents through morphometric analysis. This is my first time using this kind of software. I am been reading some information in internet and I was looking for MorhoJ sotware but it is no available on internret anymore. Could you recommendme another one. Thanks

I have a drainage network polyline feature the course of which is structurally controlled. To determine the dominant direction trends of the streams I need to plot a rose diagram. However, I am unable to derive directions of these polylines. There are 3500+ streams present in my AOI.

I wish to conduct a study involving the geometric morphometric analysis of entorhinal cortex and hippocampal shape variations. As I hope to acquire MRI scans from online databases, I would just like to ask if there are any recommended MRI parameters that accurately capture the overall shape of these two structures. For example: would it be more ideal to gather T1-weighted or T2-weighted images, and what imaging plane (e.g. axial, sagittal, coronal) would be optimal for viewing these structures?

Dear all:

Greetings,

I'm asking this question to look for experts' advice for our thesis. We are looking to study x-ray images of human hands with and without osteoarthritis, and use geometric morphometrics as a tool in trying to find something new out of it (i.e. possibly quantify the differences between the severity levels of osteoarthritis based on the alterations in the bone/joint structure). In line with this, we would want to hear your thoughts regarding what specific bones/joints are best to focus on in carrying out the study. Your answer/idea regarding this matter would be much appreciated.

Thank you and God bless!

there are many papers using channel classification as single phase and two phase but i am not able to find the definition of the both type of channels. Most of such studies used reference as Brice classification, but i am not able to find the that paper.

I need to do a morphometric analysis of a watershed using ARC GIS software. However I am unable to find the morphometric toolbox to perform the analysis. Please advice where can I get the tool box from??

I am trying to determine how the facial deformity progresses in juvenile cisco

*over time*in various treatment groups using geometric morphometrics. Our first sample (taken 1 week after being put into their treatments for acclimation), had significant differences in shape. We want to correct for these significant differences by compiling all our samples (weeks 1-8) into one PC analysis and determining the PC score for our first-week treatment groups. Could we then use these week 1 PC scores as a covariate for a Procrustes analysis on a week-by-week basis? We are already tried to correct for differences using length, but that didn't work. Could the PC score realistically work as a covariate? If not, are there any other ways to correct for differences in that first week?Using ArcMap, Archydro or any other application package

In first order watersheds, can the shape index (Ff=A/Lb²) and the elongation ratio (Er=(2/Lb)*(A/Pi^0.5) return values >= 1?

Working with large watersheds, these values tend to be < or = unity. However, working on small hydrographic basins (1st order) I find some volers above the unit.

Is there any convention that the values of Ff and Er cannot exceed unity?

Thanks.

It has been noted in many researches that gully headcuts are main drivers of gully erosion and upstream migration. Most commonly used definitions describe gully headcut as near-vertical step at which most intense erosion occurs, e.g.:

**Rengers and Tucker, 2014.**-

*Headcuts are near‐vertical steps that erode the valley network by migrating upstream over time (Bull and Kirkby ,*

*2002*

*) and add mobile sediment to gully channels downstream (Tucker et al. ,*

*2006*

*).*

**Vanmaercke et al., 2016.**-

*A gully headcut is a natural, nearly vertical drop in gully channel-bed elevation (Poesen et al., 2003).*

Since these are predominately descriptive definitions, I would like to know is it possible to delineate quantifiable definition of a gully headut ?

Such definition would be based on measurements that could be extracted from high-resolution DEM (e.g. required slope angle; headcut horizontal length; headcut height…), rather than on the descriptive, non-quantifiable information.

I understand that such definition would vary depending on local terrain characteristics and characteristics of local gully erosion predisposing factors. But even general quantifiable definition would be very helpful for detection of gully headcuts.

In the attachment is large gully headcut from my research study area, located at Pag Island, Croatia. Within Pag Island large number of small headcuts can be found, that are less distinguishable then the one in this picture.

I am working with dem images for morphometric analysis. I am using topotoolbox for Ksn calculation. In my study area there is several dams in a single river and when I am using topotoolbox, it is taking the stream reach between two dams as the longest river stretch resulting in an erroneous result. I wan to effect of the dam from the dem for further analysis.

To compare morphometric data of two or more bees.

Using TPS Series of Prof. James Rohlf for resolving problems related to morphology is a common technique in academia. I already used this series in previous successful works, however I do not remember how we can produce NTS files for Thin Plate Spline analysis. I appreciate any help from experts.

Hi,

I am creating stream orders for morphometric analysis.

At what threshold will we write to the "Con" tool when creating a stream? (Example "Value > 100").

As known, this affects all accounts as it affects the number of Strahler stream orders.

How can I determine the threshold that I am going to write in the "Con" tool and that is suitable for my basin?

Best wishes, Ahmet.

I'm using MorphoJ, and the software gives me the eigen values but it doesn't give you the eigen vector. For each landmark it gives you a x and y value for each PC but I'm looking for a value that showesme which ones vary more. Is there any command on the software to see this?

I have cell images from Confocal Microscopy and want to do Nuclear Morphometric Analysis. Can anybody share a detailed protocol to do so?

Thanks in advance.

Swarnali

SL gradient index = (change in elevation/change in length of river)*L(river length), how can we calculate the L values?

After flow accumulation analysis, what will be the criteria to define the threshold to extract the stream network from DEM? As stream threshold value will influence the extracted drainage networks and stream order level and thus the basin morphometric analysis.

I'm testing different methods of calculating missing landmarks for geomorphic morphometric analysis of rodent footprints. After an initial literature review, I came across Bayesian PCA (BPCA) and a least squares regression (LSQR). After calculating these, in R using the PCA wrapper functions in pcaMethods package for BPCA and the best.reg() for LSQR, I am left with a 2D matrix (I believe nxm), but require a 3D array (pxkxn) for all Procrustes analysis. Unfortunately I am struggling to do this step, any advice?

Note: I've tried arrayspecs() in geomorph and vecx() in Morpho. vecX() produced a matrix, but then Procrustes analysis failed to work due to an error with pcAlign.

I have exactly the same question as the person who is asking in this link:

All I know is that the number of rows and columns in this matrix are double the number of landmarks because one of them (either rows or columns) represents the variables, that is, the x- and y- coordinates of the landmarks within the tps format. The main issue with interpreting this matrix would be figuring out how does R arrange the variables. I'm guessing this would be 1x, 1y, 2x, 2y, 3x, 3y,...

Also this does not explain why columns and rows are BOTH double the number of landmarks. I'm (again) guessing that columns are (maybe) representing variables, and rows are (maybe) representing a number of principal components arranged by variation percentage explained, set by default to be the same as variables.

I am working on fish geometric morphometrics using truss network. I have fishes of one species from two sites. I have applied Principal component analysis PCA on the data to extract components. What other statistical tests can be applied on data for comparison of fish morphometrics of two sites. Moreover If I apply other tests, should I apply these tests only on principal components extracted by PCA or on all data. Thank you in anticipation!

**Trust network measurement in fish involves anaesthetizing with benzocaine (ethyl-p-amino-benzoate) [26] before being weighed and measured Figure 1. The first step is to take and record the standard length (LS), post-orbital length (LPO) and maximum body width of the fish. Standard length will be taken from the tip of the upper jaw to the base of the caudal peduncle. Then a truss network is constructed between landmark points, homologous throughout the population, chosen because they describe the major features of the fish (Figure 1). The landmarks were linked closely to the skeletal structure of fish and were easily observed by eye.**

I'm looking for any studies that have used or developed landmarks on the whole body of any species of rhino. I have lateral and posterior view photographs from 6 rhinos (both sexes) over 18 months and would like to assess how their body shape has changed during this time.

Dear All,

I performed an immunohistochemistry in mouse brain tissue looking for astrocytes (stained for GFAP). Now, I would like to evaluate any morphologic change between the different groups I am comparing.

I do know it would be possible with Fiji/ImageJ through Sholl analysis and I did read several papers suggesting so, but I do not understand what I am actually measuring.

Do you have any idea, protocol, suggestion on how to do the image analysis using either ImageJ or another software (I'd prefer an open-source software).

Thank you,

Salvo.

I tried to use Morphologika to format my data, but even the sample data set won't work (a message pops up "error in data") and the new software, EVAN toolbox, costs money to use. I want to be able to look at vectors to examine sex differences in shape to determine the severity and directionality of the sex variation at each landmark. But whenever I attempt to do the compare vector analyses, it says that there isn't enough appropriate data to analyze. Does anyone have this problem? Maybe there's a better software that I can use to study vectors/angles? Any suggestions?

A little nerdy game:

The attached document shows a selection of suture lines arranged by their values on the 1st three axes of a geometric morphometric analysis on the external suture line of Changhsingian (Late Permian) to Smithian (Early Triassic) ammonoids. This analysis follows strictly the method of Allen (2007), using Windowed, Short-Time Fourier Transform. Since this method uses a mean value between different windows, it cannot be reversed like classical fourier analysis where a shape corresponding to fixed values can be calculate in order to know what do the axes correspond to in term of morphological variation. So I made this diagram to try to decipher it. I of course already have some ideas, but would be curious what others would say about it!

Please Let me clarify my doubt how to calculate Basin length and Perimeter through ArcGIS software for Morphometric Analysis of River Basin.

Thank you in Advance.

In the attached file you can find picture with

**13 LM**as my suggestion. Many thanks in advance for all comments!I am exporting a phylogenetic tree with bootstrap from MEGA and I need the following NEXUS format to integrate the phylo data into a geometric morphometric analysis in MorphoJ software.

The following data is an example that can be imported into MorphoJ:

#NEXUS

BEGIN TREES;

Title Imported_trees;

LINK Taxa = Taxa;

TRANSLATE

1 fruticosa,

2 parvifolia,

3 palustris,

4 neumaniana,

5 crantzii,

6 pensylvani,

7 atro_Yellow,

8 atro_Gibson,

9 thurberi,

10 flabellifo,

11 aurea,

12 pimpinello,

13 argentea,

14 saundersia,

15 hirta,

16 heptaphyll,

17 thuringiac,

18 goldbachii,

19 recta,

20 dickinsii,

21 Rosa;

TREE 'Imported tree 0+' = ((((1,2),3),(((4,5),(((6,(7,8)),(9,10)),(((11,12),(((13,14),15),16)),((17,18),19)))),20)),21);

TREE 'Imported tree 1+' = (((((14,6),((16,(15,(((5,(13,4)),((7,8),(9,10))),((17,18),19)))),(11,12))),20),(3,(1,2))),21);

TREE 'Imported tree 2+' = ((((1,2),3),(20,(13,(((14,(7,8)),(10,9)),(((4,5),((17,18),19)),(((11,12),6),(15,16))))))),21);

TREE 'Imported tree 3+' = (((3,(1,2)),(20,(((14,6),(((13,(12,11)),((16,((7,8),((4,(17,18)),19))),15)),(5,10))),9))),21);

TREE 'Imported tree 4+' = ((((1,2),3),((9,(((10,5),((19,((((4,13),16),15),(17,18))),(12,((7,8),11)))),(14,6))),20)),21);

TREE 'Imported tree 5+' = (((3,(1,2)),(((12,11),((((((13,((5,4),((17,18),(15,16)))),19),(14,(7,8))),10),9),6)),20)),21);

TREE 'Imported tree 6+' = (((20,(((10,((14,(7,8)),6)),9),(((11,12),(15,(16,13))),(19,(4,(5,(17,18))))))),(3,(1,2))),21);

[etc.]

END;

Would I have to use a different software (e.g. Mesquite) to create such a format?

Many thanks,

Juniper

I ´ve recently submitted a paper dealing with Geometric Morphometrics in plants. I used PCA in order to show variation as a previous step to proceed with statistical tests (Procrustes ANOVA, DAs, etc). One reviewer asked us:

"

*Are there any reasons why PCA analysis limitations (use of variance, linear projection) should not considered important in this analysis ? This kind of question should be addressed, because they are at the heart of any interesting question anyone would ask in performing such analysis : the relations between morphogenesis and variance. It seems unclear to me, after having read the article, why kernel PCA should not be privileged versus PCA*"As I have never read about kernel PCA in GM papers, I want to know if it should be mandatory to compare it with regular PCA in these kind of papers. If not, what do you think the best response should be?

Thanks you all!

I am working on Insects (Micro-hymenopterans) and trying to differentiate the two species of the same genus using morphometric as a tool. I found that Geometric Morphometric could be used for the objective proposed. So where can I learn this tool.

Please help in this regard.

Thanks in Advance.

I have done morphometric analysis of watershed ,which gives bifurcation ratio as 4.57, elongation ratio is 0.57,shows that this watershed is elongated and peak flow is extended .(it will come early?)

means,if normal watershed what ever peak flow occur ,in elongated watershed ,the peak flow is some what in later stage

is it so!

is anyone has books related to river morphometry(specially for drainage of river is concern),I have refered but few things came across!

Concerning methods of rodents measurements: weight, sex-ratio morphologies and craniometrics.

regarding prioritization of watershed and land use pattern

I want to apply a PCA analysis with missing values for a morphological analysis using bones. Any method with missing at random (MAR) values?

Hi again

I'm dealing with some morphometric data, and I have the doubt if variation in indexes can be due to clinal variation or to separated species.

Could you recommend any paper on the subject?

Thanks a lot!

Taking photomicrographs of wings for further geometric morphometric analysis.

Wings are mounted on microscopic slides with BERLESE mounting medium

method of taking photos in geometric morphometric studies

I'm not really sure whether this diatoms are from genus

*Coscinodiscus*or*Actinocyclus*. Can anyone help me with this identification? Thank you.We are working with some fossil cones and would like to identify the species, and its closest extant relative, if possible. Is there a method to determine conifer species on the basis of cone morphology and histology?

I've found a number of tutorials on line but none of them describe step by step how to go about doing this. Any help would be appreciated. I'm including two of the images that I'm trying to process.

Handling out of vocabulary words in morph analysis

Dear all,

I am working on morphometry of a beetle community (cca. 30 species) belonging to three distinct families, but some of them clearly showing same way of life. We would like to detect which of 25+ morphological characters (mostly lenghts of different body parts, including legs) can be attributed to convergent evolution (i.e. in species belonging to different families but showing same way of life), and divergences (i.e. in species belonging to the same family but evolved differently, accordingly to their different ways of life).

Is there any explicit test for showing that? Is this possible to test without molecular data? (we know there are three distinct molecular groups, but we do not have our own molecular data)

I would be very happy to receive suggestions of any kind.

All the best,

Jure

when I use the "Residuals/Predicted Values From Other Regression" function in MorphoJ, I can't found the independent variables "data matric" and "variables"in the drop-down menus. Is there something wrong during my opretion? Thanks

I am using 2 curves with 15 evenly-spaced semilandmarks to capture the lateral morphology of a very disparate sample of bird beaks. The 2 curves are constraining by 3 regular lndmks forming a triangle, the tip-of-the-beak one constrains both curves. When I slide the smlndmks in tpsRelw with min-dsquare slide method most of the smlndmks collapse in a very narrow section of both 2 curves in the Procrustes superimposition. Minimum bending energy slide method does not affect the superimposition in this way, but I need to use min-dsquare. Any idea what is going on here? Is it a bug of the program or is a proper statistical effect?

It is possible that these molars have malformations? Or they were just growing in a wrong way ?

I identify them as

*Mammuthus primigenius?*and the black spots represents**burning traces**. Probably found near a paleolithic site.I am an MSc student and for my final thesis project I had set out to perform a comparative morphometric analysis between three avian families. My own dataset of one family returned results, however the dataset I was sent to perform the comparison with is unusable (none of us can work out why or how to fix this). Sadly, I do not have time to repeat any procedure and am having to settle with extremely restricted, unoriginal results. I have PCA results for all datasets, but as I am unable to run the datasets together, I cannot directly compare them.

I am struggling to write my report as no direct comparison between the groups was possible and as such I have no overall results. Is there a 'way' I can still write a decent report?

I just started working with Geometric Morphometrics. I am using the landmark method to test shape variation in gastropod shells. But with very few homologous landmarks, I also decided to use semi landmarks to capture shell shape. Do I have to digitize semi landmarks as curves? The outlines of my specimens are quite irregular and damaged

from srtm data i am getting stream network. in which number of stream are not correct when manually counting. how can i solve this problem?

1 st image showing wrong and 2nd image denotes correct. i am getting problem in 1 st image.

Does anyone know of any (published) 2D geometric morphometric analyses that have used coordinate data that is scaled and transformed, but

**not rotated**?program to do geometric morphometric .

Thanks in advance.

Hi, I've wrote a program that calculates the normalized elliptic Fourier coefficients of a closed polyline. I used it for a set of polylines and found that some of the polylines differ in orientation (one part oriented CW, other CCW).

I found NEFD of all polylines and kept them. After that I want to make all polylines oriented CCW, and calculate new set of NEFD. Will the values of NEFD of the polylines, that are previously oriented CW, changed or not?

I have digitized outlines of some exemplars and want to prove that they are belong to two or more different species or constitute one species.

I could not place landmarks on outlines therefore decide to use Elliptic Fourier transform. I found harmonics of EFT but don't exactly know what to do next.

Which method do I need to use to distinguish similar groups through my outlines using EFT harmonics?

I am interested in determining sex in Cape Hare from a collection of mandible.

I have not succeeded in finding anything in the literature. I do however know that sex can be determined via mandibular diastema length in some species of rabbits.

I am starting a plant morphometric study and have got a lot of inspiration from Julien Claude's (2008) book. It uses the "Rmorph" package quite much, but I failed to find its copy on CRAN pages. I would be grateful if someone could send me a copy of this package or give a link where to find it.

Hello everyone, I'm trying to make a morphometric analysis on Dakhla basin in Egypt using Arc-map 10.2, well I started to delineate the basin of Dakhla in this order: 1- Filling stage, 2- Flow Direction stage, 3- Flow Accumulation stage, after the 3rd stage is done this drainage pattern appeared with the shape presented in the attached file, I couldn't interpret what is the wrong with those straight dashed lines presented in the center of the basin. Is it an error with the DEM data, or is it an error with handling the data? I don't know, so please if anybody could help me with this, I'll be appreciated. Thanks in advance.

Dear scholars,

I am looking for a paper in which the morphometric analysis or measurements of male and female genitalic attributes have been measured/compared by considering principal component analysis for scatter plot analysis.

I am dealing with populations of certain taxa which are occurring in closely related mountain landscapes.

Critiques and suggestions are welcome.

Taking a typical claw for example, the tip of the claw is easy to determined while there is hard to find other landmarks. And there must enough (and not too many ) landmarks for geometric morphometrics analysis.

How should we do a morphometric analysis of a drainage system?

Has anyone used stationary 3D cameras to reconstruct mesh images of moving objects, and derived morphometrics from them?

I'm trying to derive the Nuclear Area Factor index as defined by Mark A DeCoster in 2007 (paper attached).

I've been thinking that morphometric analysis of the nucleus is going to be problematic if nuclear area is merely going to be a function of how deep we cut into the cell. Thus we'd have nuclei of varying sizes but they wouldn't reflect the actual size of the nucleus at its maximum girth but rather just the planar depth of cutting.

My impression is that in monolayer cultures visualised by confocal microscopy in conjunction with nuclear-specific dyes, you *do* get an impression of the full nucleus, whereas images resulting from microtomy have this issue of the planar depth variable. Incidentally some papers I have encountered have ignored this issue and gone ahead with morphometric analysis on nuclei in histological sections.

Do you have any suggestions on how to deal with this? I cannot do another experiment. I can only make use of the ultramicrotomy sections I have, in which cells are cut at a variable depth of plane through fixed liver.

I'm analyzing certain structures in annelids through scanning electron microscopy, and I would like to obtain morphometric data for comparisons.

Could you recommend me free software to do it?

Some features I would appreciate are easy scaling of different images and an intuitive interface.

Thanks.

Let's say, I have landmark data for three different structures (A,B,C; e.g. three different teeth along the tooth row or fingers on one hand) for one sample of specimens and I would like to know whether structure A is stronger integrated with structure B than structure B with structure C. Is it possible to compare RV-coefficients calculated for the A-B and B-C interaction and would the resulting difference be valid to make inferences on relative integration strengths?

I have already asked some colleagues about the problem but got somewhat contradicting answers. So I am curious about your opinons. Maybe comparing the values would be possible under the condition of the same landmark number on all structures?!

Best regards,

Stefan

Drainage Texture ratio is Morphometric analysis parameter used to estimate erodibility.

I'm doing morphometric analysis of splenocytes and from the data that I have processed up to now I see quite strange numbers for Feret diameter and I wanna know how big are splenocytes?

Some publications indicate size that range from 8-12 um (peripheral blood Ly) while altered cells (enlarged) above 12 um. How ever other textbook data are that the size of Ly - normal - is fro 8-25 um! This "subtle" differences are the point of my work/analysis can you help me with some suggestions/references?

I have a plan to conduct a study about the different cultivar of cassava found in Romblon. And, to determine the phylogenetic tree of cultivar, what kind of morphometric analysis should I use?

Thank you. :)

Except resolution of 30 meters, How far is the ASTER DEM accurate for the mapping, for morphometric analysis or any other purpose of study, over SRTM DEM?

I want to measure aorta diameter but the shape is not find round.