As a preface, let me acknowledge that this blog has been perhaps a bit too preoccupied with statistical genetics in the past month. Looking at the several posts I have partially completed it is likely to remain so through the summer.

**Sequencing and GWAS
**

Nature Genetics has in the past few months returned to its old position as a clearinghouse for GWAS studies, the author lists for which steadily increase as more and more groups contribute data. (Can we start putting the author list in the supplemental material? It’s getting hard to navigate the webpage…). GWAS studies are nice, we sort of know how to do them. I have a bit of a beef with the push-button approach, I think GWAS studies and in particular meta/mega analyses are failing to deliver as much information as they should, and I’ll come back to this at the end of the section. But GWAS always have the complexity that they are *ascertained*. We didn’t find a variant of effect T. Does one exist? Can’t say, we could only query these X variants.

Sequencing is fundamentally different: whatever effects are driving the phenotype in your sample are present in your sample. Yet the treatment of (particularly exome) sequencing studies is “like a GWAS just with more variants,” with no real use of the fact that variants are discovered rather than ascertained. It’s the same kind of “where is the signal I can find?” question. But really, the trait is heritable, the signal is there, somewhere, and it’s not just about the signals you can find, it’s also about *the places you can rule out*. This applies to GWAS as well: you genotyped 50,000 samples and have 12 hits. Surely you can say *something* else about the other 199,988 sites on the chip, right? Or at least about a large fraction of them?

And once you go from ascertainment to discovery, you can rule out whole portions of the variant space. Things like “No variant in SLC28A1 of frequency > 5% has an odds ratio >1.2”. With that said do I still care about SLC28A1 as a candidate gene for my trait? What if the best I can do for ADAM28 is to say no such variant exits with OR>2.6, but may exist for OR<2.6? Now how much do I care about SLC28A1? You can make such statements having performed sequencing, not only can you produce a list of things where signal *definitely is*, but you can segment the rest into a list of things where the signal *definitely is not*, and those where signal *might* be.

In some sense GWAS studies dropped the ball here too, but the analysis is far more subtle. One of the big uses of modern sequencing data is *fine mapping*: so, at a GWAS locus, trying to identify rare variants that might be generating a “synthetic association”. For instance, if several rare variants of large effect happen to be partially tagged by the GWAS hit. However, given the observed variance explained, there’s then a posterior estimate of frequency, effect, r^2, and number of variants that could be consistent with the observed signal. Carefully working out what *could* be there, and what’s *expected* to be there would provide not only intuition about what to expect when performing fine mapping, but a good and precise framework for thinking about synthetic associations. While the idea that most GWAS loci are so explained has been vigorously rebutted, it is done so at the level of bulk statistics, with little attention paid to *if* there is a synthetic association *here*, what must such variants look like?

The point is ultimately there’s a lot more information than *what things have p<5e-7*, it just needs to be extracted.

**Bounding Variant Effects**

The 0th-order question is “What variants do I have with *p<5e-7*“. If the answer isn’t the empty set, the 1st-order question is “What genes are near these variants?” and the 2nd-order question is “What biological processes are these genes involved in?”. If the answer is the empty set, the 1st-order question is “What gene burden tests* *are at the top of the quantile-quantile plot?”, the 2nd-order question is “What biological processes are these genes involved in?”

But, as I mentioned in the previous section, you’ve seen these variants enough times (especially now with tens- to hundreds of thousands of samples) to know whether or not they’re interesting. The first-order question ought to be “What variants can I *rule out* as contributing substantially to disease heritability [under an additive model]?” Ideally, this would be effectively everything you looked at. In practice, you looked at 100,000-10,000,000 variants, and some of those will look nominally associated by chance, and these will be difficult to rule out.

Alright so what am I talking about? Here’s the model:

where *X* are the genotypes at a site, and is the *i*th covariate. It is the property of maximum likelihood that, given the data, the coefficient vector ). Denoting the variance-covariance matrix as then . This posterior distribution on the estimate allows more than the basic test of , one can also ask for what values we have , . This is just a *p*-ile confidence interval, but if *p<1e-6* and is small with , then this particular variant is very unlikely to be causal, given these bounds, in that the probability of a large beta is very low.

Now there are two problems with “just using a confidence interval.” First, because the likelihood is heavily dependent on the number of observations of nonzero , the variance of the estimator is an increasing function of the minor allele frequency. That is, rare variants are harder to bound. Secondly, the disease heritability *conferred* by a variant of given effect increases with allele frequency. That is, a 1.4 odds ratio variant at 0.5% frequency is, in and of itself, contributing very little to the prevalence of the disease. By contrast, a 1.4 odds ratio variant at 30% frequency is contributing a large fraction of disease heritability.

Both problems can be resolved by taking a heritability view, that is, asking instead the question for what proportion of phenotypic variance we have

In other words, say you observe a variant with , at a frequency of . You then ask, what is the probability that this variant explains 1% of the total prevalence (on a liability scale) of my disease, given this observation? Well, that’s simple, given the prevalence and frequency, one can simply ask for which two values of beta, , the risk-effect and protective-effect values, is the proportion of phenotypic variance 1%. Then, given those values, use the posterior distribution to ask what is ?

Note that the *frequency* of the variant does not enter into the discussion, it’s only hidden in the calculations of the phenotypic variance , and the standard deviation of the maximum likelihood statistic , thus making variants of different frequencies commensurate by placing the hypotheses on the same scale – and one growing approximately with the variance of the estimator. That is, while the variance of the estimator increases as the variant becomes rare (this is what makes bounding using a constant odds ratio difficult), so too does the effect size required to confer a given level of phenotypic variance. These phenomena tend to cancel, so on the scale of variance explained, one can bound a rare variant as well as a common variant.

In order to do this, it’s just a question of converting a given variance explained into frequency-effect pairs, and evaluating the likelihood of such pairs given the data. That is:

These terms need to be integrated over , but for whatever reason the wordpress parser doesn’t like it when I put $\int_{f_{\mathrm{true}}$ into the above equations. And in practice, for even rare variants on the order of 0.5% frequency, the posterior frequency estimate is peaked enough to be well-approximated by setting (remembering, of course, that all frequencies are population-based, so oversampling cases requires re-adjusting the observed frequency based on the prevalence of the disease: ).

The last thing to work out is the function giving the phenotypic variance explained by a variant of frequency , effect size on a dichotomous trait of a given prevalence. It’s very well known how to compute the liability-scale variance, and the inverse can be found trivially through numerical quadrature.

Liability-scale Variance: A DerivationRecent discussions have left me slightly cynical on the common intuition behind this calculation (an intuition which seems to be “cite So et al 2012”), so it may be worth stepping through the derivation. Let’s start with a disease, and a variant. The diesease is 10% prevalence, and the variant has a 10% frequency, and an odds ratio of 1.35 (so ). We begin with assuming an underlying latent variable with some distribution, and for the sake of having a pretty function, I’m going to assume a logistic. There’s also a choice between setting the mean of latent variable (over the population) to 0, and letting the threshold T adjust, or setting T=0 and letting the mean adjust. I will do the latter. So basically we have a whole population that looks like

Where the area under the curve to the right of 0 is 0.1. The population mean is exactly . So the question is: whereabouts in this liability scale do carriers fall? For that we need to know the probability of being affected by genotype state. Given the odds ratio, this is pretty easy:

There are three equations and three unknowns (plug in ), and so this system is soluble. This can be reduced easily to a single equation by plugging in, and solved by a simple root solver such as

diseaseProbs <- function(frequency,odds,prevalence) { bt <- log(odds) cPrev <- function(muR) { PR <- (1-frequency)^2*logistic(muR) PH <- 2*frequency*(1-frequency)*logistic(muR + bt) PV <- frequency^2*logistic(muR+2*bt) (PR + PH + PV - prevalence) } if ( odds > 1.05 ) { # prevalence almong reference individuals will be lower res <- uniroot(cPrev,lower=invlogistic(0.001),upper=invlogistic(prevalence)) } else if ( odds < 0.95 ){ # prevalence among reference individuals will be higher res <- uniroot(cPrev,lower=invlogistic(prevalence),upper=invlogistic(0.99)) } else { upr <- invlogistic(min(1.2*prevalence,1-1e-5)) res <- uniroot(cprev,lower=invlogistic(0.8*prevalence),upper=upr) } muR <- res$root return(c(logistic(muR),logistic(muR+bt),logistic(muR+2*bt))) }So what does this give us? Well it gives us the Logistic Mixture comprising the population distribution

Where I haven’t beenentirelyfaithful to the densities (I’m scaling by the square root of the frequencies of the genotypes). Point is, you can see the shift in the distributions by genotype, and consequently the proportion of the distribution curves falling above theT=0threshold. At this point, we can just plug into the formula for variance: to getOf course throughout we have assumed a standard Logistically distributed error term. Thus the variance of the means have to be given to us as a proportion of the total variance, or in other words . In this case, the variance of the logistic is , and thus we set

So roughly 0.5% of the liability-scale variance. Note that using the standard method of replacing the logistic distribution with a standard normal, and using results in an estimate of 0.44%. These differences increase with the variance explained, so that for a 20% frequency variant with OR=2.35, such a variant explains roughly 6% in the logistic model, but 5% variance in the normal model.

Assuming the posterior distribution of *f* is sufficiently peaked, doing a GWAS-like bounding scan is fairly simple. For each variant, calculate

via . Then, using the posterior distribution of calculate

.

**Multiple Test Considerations**

Where does the figure 1*e-7* come from for GWAS studies? Or *5e-8* for sequencing studies? (And I mean conceptually, not here here or here). The basic logic is that of the *look elsewhere effect*:

1) Most variants are null

2) There are lots and lots of variants

3) There are many chances to see spurious nominal associations

4) Must set the threshold fairly high to achieve a study-wide Type-I error of 5%.

The fundamental consideration is (1), and there’s a significant asymmetry between a GWAS scan to reject , and a bounding scan to reject . If most variants are null under a GWAS scan and ought not to be rejected, then almost no variants are null under a bounding scan, so most ought to be rejected. That is: Type-I error is bad in a GWAS scan (you find something that’s not real), and you’re willing to pay a cost of Type-II error (you don’t find something that is real). By contrast, for a bounding scan, Type-II error is bad (you put bounds on a truly causal variant), while Type-I error is less of an issue (you fail to bound a noncausal variant).

Thus, previous conversations about multiple testing are not applicable, and one shouldn’t blindly apply *p=5e-7*. Instead, bounding requires a good deal more thought (or assumption) about the architecture of the trait. I would articulate the logic as

1)

Causalvariants are null2)

Fewvariants are null3) There are lots and lots of variants

4) There are many chances to falsely

acceptthe null5) We can be rather aggressive on the rejection

p-value

It’s worth articulating why we don’t care all that much about false rejects: rejecting the null *falsely* simply includes a non-causal variant in a set of variants that should be enriched for causal variants. Variants for which the correct test is to reject . And it’s absolutely possible for a variant to be such that (e.g. have a large enough standard deviation that) is rejected, as well as is rejected. On the other hand, *failing* to reject the null on a *causal* variant means we can potentially exclude it from follow-up analyses, a situation we want to avoid.

So there’s a different balance between Type-I and Type-II error here, with Type-II being the one we’re concerned about. But of the two, Type-II error is the trickier, because it’s fundamentally a question of power. The idea is, well, we don’t want to accidentally exclude things. But you have to be a little bit more specific: what kind of things do you not want to exclude? You can be sensitive to all causal variants (even those with OR=1+1e-5) by rejecting everything. So being more reasonable, you might say “I want *no* false acceptances of any variant with >0.5% of phenotypic variance explained”. Given that statement, you can calculate the power , and that’s the power at the nominal threshold . (See previous posts regarding calculation of power). But of course, that’s not *quite* enough, because you also need to know the number of chances to falsely accept such a variant, which is, of course, begging the question (if you knew how many causal variants there were, why bother with the exercise?). One way to get around this is to simply assume an unreasonable number of causal variants of such variance explained, and then use the same machinery as Bonferroni correction and set

**Bounding Variance Explained at a Locus**

Alright, so that all was about variants you know about. But what if you care about a specific locus, like a gene, and want to know if variants within that gene contribute substantially to heritability. There are several different questions to articulate, but for now let’s stick with the single-variant approach. We know, from the previous section, how to answer “what is the probability that one of the variants in this locus explains a *T* or greater portion of phenotypic variance?”

But now we’ll take a step into the unknown. The *second-order question*: what is the probability that *no variant exists at this locus that explains a *T *or greater proportion of phenotypic variance? *That is, given I’ve sequenced the locus and haven’t found anything, what’s the probability I could keep on sequencing and *still* wouldn’t find anything?

Well, what’s the probability the sun will rise tomorrow?

Following Laplace, 9592/9593.

Well Okay. In all seriousness, this particular question about genetic mutation has just enough structure in it to avoid the philosophical issues that arise with the Sunrise Problem (Statistics can’t prove a negative, etc). Specifically: we rely on the fact that we’ve done sequencing. This means that we have *not* failed to query sites for mutations (as in the case of genotype chips), but we have, and there were none. That is, if a variant exists, we’ve got a pretty good idea about it’s frequency. Let’s turn to Mister Bayes, and ask, given I’ve sequenced samples and seen counts of a given allele, what is the posterior density of the allele frequency? Using the Rule:

This doesn’t give a good intuitive sense of the posterior distribution other than the basic shape (gamma-like, no?). The best thing to use as a prior is the allele frequency spectrum modeled on the human bottleneck-and-superexponential-expansion (see Evans and Shvets, for instance), but I find a good intuitive model (that surprisingly works well in practice) is a truncated inverse distribution

which is distributed on the domain . This tends to overestimate allele frequency because (1) humans have grown faster than exponentially, and (2) no mutation is allowed to have a frequency lower than . You can make a Gamma distribution peak at and then go to 0, but this is a nice toy distribution. Plugging it in yields

Digression: The Probability that a Singleton is more than a SingletonThis is actually a fun little proof I have given this simplified prior. You go out and sequence

Npeople, and find a singleton you care about. What’s the probability that it’s frequency is >1/2N? I’ll prove that it’s about . What’s the denominator of the above formula when ? Plugging inAnd therefore the conditional posterior of

fcan be writtenNow we can inquire as to the probability that, given AC=1 in N people, the

truefrequency will be greater than some value . This is:Now let :

Letting yields:

Anyway, the point is you have some posterior estimate of the allele frequency spectrum given that you haven’t observed it in your sequencing. Any variant you haven’t seen yet has that posterior spectrum, and thus given a fixed proportion of phenotypic variance explained *T*, the odds ratios that such sites must have are constrained to be very large. In particular, for a 10% prevalence disease, using the above model, a 0.01% frequency variant would need an odds ratio of 400,000 to explain 1% of the liability-scale variance.

However, the fact that for such a rare variant, 1% phenotypic variance is achievable is really a breakdown of the liability-scale model. In particular, there is a ceiling on the variance explained by a mutation: a 0.01% variant that is fully penetrant only explains 0.1% of the prevalence of the trait. That is, at some point you know that any undiscovered single variant left at a locus, *even if fully penetrant*, cannot explain a significant fraction of the phenotypic variance.

On the observed (0/1) scale, such a variant of frequency *f* for a disease of prevalence *p* explains a proportion of the phenotypic variance. Transferring this to the liability scale by Dempster’s formula

Now the genetic variance on the observed scale, as the slope is at most 1. by definition, and therefore . Obviously during the conversion to the liability scale, the phenotypic variance, which appears in both denominators, will cancel. Thus we have (for prevalence ):

.

Here, so for and 8% prevalence, or 0.45% of the phenotypic variance. This is a bit of a tawdry estimate, as such a variant equipped with OR=15 explains (by the other model) 0.14% of the phenotypic variance. Nevertheless, this transformation provides a reasonable approximation to an upper bound for rarer variants (but we should still search for better corrections to it). Thus, denoting by

the function mapping a frequency to the maximum liability variance explained (e.g. that when the variant is fully penetrant), the probability that an undiscovered locus explains fraction of phenotypic variance is

where is the minimum *f* such that . Note that this expression is also , that is, bound by the posterior allele frequency distribution function evaluated at .

Now is easy to calculate, set so that

Thus given the prevalence of a disease, there is an effective upper bound on the liability-scale variance explained, given that the observed 0/1 variance cannot exceed . Here, we plot as a function of the variance explained

So let’s go to our example. Lets say you’ve got a 10% prevalence disease, and have sequenced 50,000 individuals at a locus. You want to know the probability that there’s a 1% variance explained mutation hiding somewhere in your population that you didn’t discover. Then V=1% and z = 0.17, plugging in gives . Using the posterior from above, . Here’s the kicker: Even if you’ve sequenced only 10,000 individuals. So long as you haven’t seen a variant *at all*, the chance that such variant that could explain 1% of the variance is still < 1e-11. That means that for a 1e-6 “chance” to have missed such a variant, about 100,000 such variants would have to exist in the population you sampled. At the locus you’re looking at. Quite reasonable on a genome-wide scale. Quite *unreasonable* for a gene.

In other words, if even a moderate amount of sequencing has been done on your gene, **there are no more single variants left to**** find.**** **

At least, if you care about single variants explaining a large portion of heritability. Other caveats: this assumes a *homogenous* population, it could be that there are populations where such variants do exist even at reasonable frequencies.

But wait, there’s more. Let’s say such a variant *does* exist. It’s completely penetrant, but it has a frequency of . For the sake of example, let’s say you’ve sequenced people from Dublin, and this variant at frequency *1/50,000* didn’t get into your study. Or maybe you did see a copy or two, it actually doesn’t matter. Suppose it’s causal dominant, you get the disease if you’re a carrier.

But here’s the deal. Dublin’s population is 1.3 million. Only 26 copies of this allele exist. You could sequence every person in Dublin and you wouldn’t have the power to associate it at *5e-7. *Furthermore, there are constraints beyond population size: you need to actually collect the samples, and you need to pay for the sequencing. Thus, in addition to there not being single variants of high variance explained left to find, *many of ***those things we haven’t found yet****cannot be demonstrably associated. They are un-followupable.
**

The good news is, the things we *have* found *can*. But at some point the posterior allele frequency becomes so scrunched towards 0 that (on a single-variant level at least) you just have to forget about ever following those up statistically. At that point you’ve got to break out the crispers and engineer some cells.

*Another Comment on Studies*

The preceding discussion is basically the entirety of the reason sequencing studies have really dropped the ball regarding the genetic architecture of the traits they examine. Sequencing studies are effectively mini-GWAS studies, characterized by a Q-Q plot and a multiple-testing burden higher even than that of GWAS, with no clear understanding or mention of why sequencing was undertaken in the first place. “We wanted to discover all the variants present in our gene.” Okay fine. Given you’ve done that, do we ever need to sequence that gene again? What’s the probability that we missed something there that’s explaining a large fraction of the variance of the phenotype? You don’t even need phenotyped samples to answer that question, you can just use estimates from the 1000 Genomes Project!

This is what’s frustrating: genetic studies cherry-pick one or two novel top hits, and leave everything else on the table. The process of *exclusion* has been entirely ignored. If we went back, and actually analyzed the data with a careful eye, we probably would have completely excluded hundreds, maybe thousands of loci from consideration. Certainly thousands of single variants.

**The Multi-Variant View**

My little (now reaching ~3500 words) invective, up til now, has been focused on a single-variant view of heritability (variance explained). This view, while helpful for talking about what can be true about the properties of single variants, is less helpful about talking about what can be true about *sets* of variants. Things like “variants in SOX9” (a very cool gene), or “variants in open chromatin regions near actively transcribed genes in renal cells.” This section is basically an extension of the previous discussion to groups of variants; but this alteration presents some novel challenges: the nonlinearity of heritability estimates, the large state space of multi-variant genotypes, and the statistical brutality of linkage disequilibrium.

*Expanded Multivariant View*

Now, rather than having a single variant , one has multiple variants , with frequencies , and effects . It’d be very nice (and indeed, some papers do) to write

But unfortunately is linear neither in nor in . First, is decidedly nonlinear in V. Second, is quadratic in both and , and is quadratic in and inverse logistic in . So forget that sum. Instead, it is straightforward to extend from a genotype to a multigenotype.

Multiple Variant Variance ExplainedThis follows from replacing the index (Ref,Het,Var) from the previous calculation with the multi-index ([Ref,Ref,…],…,[Var,Var…]). For convenience, we’ll wrap the multi-index to a univariate index , and denote by the genotype of the th variant in the th multigenotype. Then:

The frequency of multigenotype we denote by , which, in the case of full HWE with no LD is

Then using

The only issue is the sum. As in the case of power (which requires massive sums over genotype space), we can get around this more efficiently via simulation. We can exploit the fact that we can set and allow T to shift. It’s easier also to use the standard normal rather than the logistic for this (as a sum of normals is still normal) A simulation would then be:

multiVarSim <- function(frequency,odds,prevalence,logit=T,prec=0.01) { bt <- log(odds) invlink <- qnorm link <- pnorm if ( logit ) { invlink <- invlogistic link <- logistic } # this part can be done with replicate, but I enjoy vector algebra nRuns = round(10/prec,1) drawGeno <- function(f) { rbinom(nRuns,size=2,prob=f) } S <- as.matrix(sapply(frequency,drawGeno)) %*% as.matrix(bt) # now u_0 needs to be such that the total prevalence is prevalence # this means that sum(link(S+u0))/length(S) = prevalence cPrev <- function(muR) { sum(link(S+muR))/length(S) - prevalence } res <- uniroot(cPrev,lower=invlink(0.001),upper=invlink(0.999)) muR <- res$root S <- S + muR # now remember we care about the variance around muP sum((S - invlink(prevalence))^2)/length(S) }

The only trouble with using the multi-genotype approach is that the state space is huge: for *n* variants, there are genotype assignments, so forget about looking at 15 variants, let alone hundreds. As mentioned, simulation can overcome this setback, but simulations themselves are expensive.

But even beyond that, when one moves from *calculating* variance explained by several variants to *bounding* variance explained by several variants, the numerics becomes challenging. In particular, one moves from a one-parameter posterior distribution (two-parameter if taking uncertainty of the frequency into account) to an *n*-parameter distribution (2*n* if accounting for frequency uncertainties), with non-axial limits of integration. That is, while for a single variant

produced a single point , for multiple variants

produces a curve:

That is, you can reduce the effect size of one variant, while increasing the effect size of another variant to compensate. This is a situation where one might use Monte Carlo simulation to evaluate the integral: but note that for each point of the Monte Carlo, a simulation needs to be run to get the variance explained. This is terribly costly.

Another way to *bound* the probability is to solve the constrained optimization

The likelihood is (luckily) convex, so this isn’t excessively difficult to do. Let be that maximal p-value. Then

Nevertheless, even this is very costly. For every evaluation of by the optimizer, a simulation needs to be done to estimate its value; even this approximation is really expensive. In addition, LD generates additional difficulty: Haplotypes need to be simulated rather than genotypes (in order to estimate ), and if the were fit independently rather than jointly, both the mean and the the variance-covariance matrix of the posterior distribution need to be adjusted to account for LD.

This, combined with the problem of estimating effect sizes from low-frequency variants, leads to methods of collapsing variants across a locus into scores.

*Collapsed Multivariant View*

Collapsing is a means of re-writing the multiple-variant joint model. Instead of

we write as a matrix and have

Where is constant and for the calculation of variance explained, . The point is itself is not fit, but only the coefficient which, when , . Collapsing tests (“burden” tests) don’t have access to , but instead try to “guess” using some function of the variant properties (like frequency, estimated deleteriousness, and so on).

One thing immediately recognizable is that if and then the formula for

So in other words, the score vector is an estimate of the latent liability variable for all of the samples. This allows the following simplifying assumption:

for some and . This is sort of more-or-less true as are a sequence of binom(2,f_i) variables, and so a weighted sum should be approximately binomial, ergo approximately normal. Therefore, for a given :

Variance Explained by a Collapse TestWe have . Here we use a normal latent model because it plays nicely with the assumption that is normal. Then

As before, can be found by numerical quadrature. Then

and .

Note that this can be simplifed by talking about a variable , but it’s nice to clarify what does (which is

scalethe proposed score distribution so it aligns with the underlying liability explained).To use this as an approximation to the variance explained by multiple variants, note that is a linear function, and therefore (assuming no LD and no relatives):

Expressions which can be adapted for the presence of LD and relatedness.

This gives and

For an evaluation of “true” variance explained, set . To identify the variance explained by a (possibly wrong) guess , replace all instances of with .

Using a collapsed estimate of the variance explained has the benefit of being univariate. That is, for a given (either guess or true) , then , and thus it has a univariate inverse: . This enables the bounding of a locus or burden test through the posterior distribution of the parameter , just as we did with a univariate in the first section. That is, given (and assuming we don’t need to integrate over the posterior of which is fairly painful in this case), to query the probability of a locus explaining Q or more of the phenotypic variance:

Calculate

Calculate

*However* (!), the obvious approach (set ) where was fit jointly by MLE has the drawback that is inflated due to overfitting. In fact, in such cases always. One way around this is to use estimated from a different source. Alternatively let be a *hypothesized* score (e.g. a burden weight vector). It’s important to note in the latter case that bounding variance explained by a burden test does not bound the maximum possible variance explained at a locus, it bounds the variance explained by the hypothesized . The variants at the locus could still contribute substantially to trait heritability, just simply in a direction orthogonal to (of which there are ).

**Surely the many things I haven’t seen explain lots of heritability in bulk, right? Right?**

The last thing to check off with the locus view now that we can bound variance explained by (1) multiple variants and (2) burden scores, is the probability that at said locus, there may exist *many* variants that for whatever reason were not discovered, but that *in aggregate* explain a large proportion of heritability. As in the single-variant case, you can put bounds on these things, but more importantly, consider what it means to want the answer to this question (as anything more than an exercise in probability).

It’s been a long post. Let’s take a step back.

Human medical genetics is science with an ulterior motive, where the search for additional causal variants is often explicitly motivated by the statement that new loci provide new targets for pharmaceutical treatments. And, by and large, this is a positive force for the advancement of disease biology and the understanding of human disease. At the same time, I believe that this Quest For Treatment, combined with the increasing focus on impact explains a wide variety of phenomena in the medical and statistical genetics community, from hasty and opportunistic study design to the ENCODE backlash (and the resurgence of Big vs Small science).

Sequencing is a good example. Initial GWAS studies provided a good number of variants and loci with strong disease associations. The bulk of these associations, however, were not immediately interpretable: falling between genes (unclear which protein might be affected), or in a noncoding part of a gene (so not obviously gain- or loss-of-function). In part because of the difficulty of interpreting biological effects, and in part because of the worry of a winner’s-curse effect, we started (rather uncritically) to refer to these variants as “tag-SNPs”, hypothesizing that the true causal variant was unlikely to have been ascertained, as the bulk of human variation is uncommon to rare and not present in the set of imputed loci, and furthermore is likely to have an interpretable biological impact. In retrospect, it seems rather clear that common variants which maintain their association across populations are unlikely to be tagging a “true causal variant” of lower frequency which is far more likely to be absent in other populations. (That said, one proof of an elegant experiment that the results reveal something that is obvious in retrospect)

I think the shift from array-based GWAS towards sequencing-based studies reflects both the optimism of researchers, and the entrance of impact as a scientific consideration. Should loss-of-function mutations underlie even 10% of the GWAS hits, we’d then have generated a respectable number of druggable targets, opening up new paths towards treatment. Even having seen the (largely negative) results, even as I* *agree with Kirschner’s characteriztion of impact, I *still* think these were the right designs to follow up on the results of GWAS.

But what it *does* mean is that these studies weren’t designed to add to the knowledge of genetic architecture. Why didn’t we only sequence cases that had none (or very few) of the known genetic risk factors? Because at the same time we wanted to see if there were un-discovered variants underlying those GWAS signals. Why do we sequence exomes instead of the regions near GWAS hits? Because not only do many GWAS loci fall near to or within genes, but also protein-coding variants, in particular loss-of-function variants, provide a very clear target for both follow-up lab experiments, and potential therapeutic developments. In addition, exome sequencing is cheaper so one can sequence more samples to find more rare variants and have more power to associate them.

So, really, the question “What’s the likelihood that there are *multiple *variants that I have *yet to ascertain* that in aggregate explain a substantial fraction of phenotypic variance?” is a continuation of this rather incautious optimism about 1) the strictly additive model holding, and 2) the infinitesmal (“Quantiative”) model not holding, to say nothing of 3) unbiased estimates of heritability. For a common disease, there are reasons on all three fronts to cautiously articulate hypotheses before going any further in the Quest for Treatment.

As mentioned in the last section on bounding the variance explained by undiscovered variants, we’ve now gotten to territory where undiscovered variants are so rare that in order to explain even 0.1% of the phenotypic variance they have to be effectively 100% penetrant. The fact that common diseases are not known to present as* Mendelian* in a substantial portion of cases, as the assumption that extremely rare (or private) variants of strong effect would suggest, indicates that such variants are not likely to account for a large fraction of disease prevalence. At the same time if these variants are not of high effect but* *together account for a large fraction of heritability, the model isn’t really distinguishable from the Quantitative model: every variant is causal, the effect size is negligible.

At the same time, the Latent Variable Model is just an approximation, and while it alleviates some of the issues of treating a binary trait as a real-valued trait (with actual values of 0 and 1), it does not eliminate them. That is, phantom epistatic effects are mitigated, but not not eliminated (because the latent variable need not be normal, though we assume so). This not only impacts estimates of heritability, but also indicates that the purely additive model is mis-aligned, and there might be a considerable gain (in terms of understanding heritability estimates) to carefully considering what happens in individuals with many genetic risk factors. At the same time, both association scans and association study experimental designs tend not to consider maternal, dominance, additive-additive, or additive-environment effects. Which seems like a more fruitful use of research money: sequencing many thousands of samples in the hope of finding a fully penetrant *f=*1/30,000 variant, or collecting a wide array of phenotypes and environmental data to better understand endogenous and exogenous disease and gene x disease covariates? Or buying a cluster (or access to Amazon’s) to scan for additive-additive effects?

In sum: none of this proves that a locus can’t harbor undiscovered variants that explain some given fraction of phenotypic variants. One can bound that probability just as in the single-variant section. But the point is: think about *why* such a bound is necessary. Is the working hypothesis to be tested *really* that there are high-penetrance near-private variants lurking in the genome? Is that really the hypothesis Occam’s Razor would suggest, given all of the observations in this post?

I, for one, don’t think so.

[…] in modeling the distributions of read counts or genotyping errors. I have written about specific cases before. The general conclusion is that likelihood-based regressions result in powerful tests with […]

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